Upload
lyngoc
View
223
Download
6
Embed Size (px)
Citation preview
Hydrogels for Regenerative Medicine:
Development and Characterization
Dissertation zur Erlangung des Doktorgrades
der Naturwissenschaften (Dr. rer. nat.)
der Fakultat fur Chemie und Pharmazie
der Universitat Regensburg
vorgelegt von
Ferdinand Paul Brandl
aus Hirschau
November 2009
Diese Doktorarbeit entstand in der Zeit von Dezember 2004 bis November 2009 am
Lehrstuhl fur Pharmazeutische Technologie der Universitat Regensburg.
Die Arbeit wurde von Prof. Dr. Achim Gopferich angeleitet.
Promotionsgesuch eingereicht am: 17.11.2009
Datum der mundlichen Prufung: 15.12.2009
Prufungsausschuss: Prof. Dr. Sigurd Elz (Vorsitzender)
Prof. Dr. Achim Gopferich (Erstgutachter)
PD Dr. Rainer Muller (Zweitgutachter)
Prof. Dr. Armin Buschauer (Drittprufer)
Meinen Eltern
in Liebe und Dankbarkeit.
Contents
Hydrogels for regenerative medicine 1
1 Introduction and goals of the thesis 3
1.1 Principles of regenerative medicine . . . . . . . . . . . . . . . . . . . 4
1.2 Biomaterials for regenerative medicine . . . . . . . . . . . . . . . . . 5
1.3 Goals of the thesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6
2 Rational design of hydrogels for tissue engineering 11
2.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13
2.2 Environmental factors as morphogenetic guides . . . . . . . . . . . . 14
2.2.1 Integrins as mechanoreceptors . . . . . . . . . . . . . . . . . . 15
2.2.2 Mechanical cues regulate cell behavior . . . . . . . . . . . . . 16
2.3 Mechanical properties of materials and their characterization . . . . . 17
2.3.1 Atomic force microscopy . . . . . . . . . . . . . . . . . . . . . 20
2.3.2 Magnetic resonance elastography . . . . . . . . . . . . . . . . 21
2.3.3 Monitoring of cellular traction forces using fluorescence reso-
nance energy transfer . . . . . . . . . . . . . . . . . . . . . . . 23
2.4 Rational design of hydrogels for tissue engineering . . . . . . . . . . . 24
2.5 Physical parameters regulate tissue development . . . . . . . . . . . . 28
2.5.1 Impact of mechanical factors on cell function and tissue mor-
phogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28
2.5.2 Influence of degradation profile on tissue formation . . . . . . 30
v
Contents
2.5.3 Cell-responsive hydrogels . . . . . . . . . . . . . . . . . . . . . 32
2.6 Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34
3 Poly(ethylene glycol) based hydrogels for intraocular applications 37
3.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39
3.2 Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . 41
3.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41
3.2.2 Synthesis of PEG-amines . . . . . . . . . . . . . . . . . . . . . 41
3.2.3 Synthesis of branched PEG-succinimidyl propionates . . . . . 42
3.2.4 Preparation and rheological characterization of hydrogels . . . 44
3.2.5 Characterization of hydrogels by NMR . . . . . . . . . . . . . 44
3.2.6 Cytotoxicity of cross-linked hydrogels . . . . . . . . . . . . . . 45
3.2.7 Release of FITC-dextrans and fluorescent nanospheres . . . . 46
3.3 Results and discussion . . . . . . . . . . . . . . . . . . . . . . . . . . 47
3.3.1 Preparation and rheological characterization of hydrogels . . . 47
3.3.2 Characterization of hydrogels by NMR . . . . . . . . . . . . . 52
3.3.3 Cytotoxicity of cross-linked hydrogels . . . . . . . . . . . . . . 53
3.3.4 Release of FITC-dextrans and fluorescent nanospheres . . . . 54
3.4 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56
4 Hydrogel-based drug delivery systems 57
4.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59
4.2 Materials and methods . . . . . . . . . . . . . . . . . . . . . . . . . . 60
4.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60
4.2.2 Synthesis of polymers . . . . . . . . . . . . . . . . . . . . . . . 61
4.2.3 Rheological characterization of hydrogels . . . . . . . . . . . . 61
4.2.4 Equilibrium swelling of hydrogels . . . . . . . . . . . . . . . . 62
4.2.5 Calculation of hydrogel network mesh size . . . . . . . . . . . 63
4.2.6 Fluorescence recovery after photobleaching (FRAP) . . . . . . 64
4.2.7 Nuclear magnetic resonance (NMR) spectroscopy . . . . . . . 66
4.2.8 Release of FITC-dextrans . . . . . . . . . . . . . . . . . . . . 66
4.3 Results and discussion . . . . . . . . . . . . . . . . . . . . . . . . . . 67
4.3.1 Physicochemical characterization of hydrogels . . . . . . . . . 67
vi
Contents
4.3.2 Estimation of diffusion coefficients . . . . . . . . . . . . . . . . 69
4.3.3 Determination of diffusion coefficients by FRAP . . . . . . . . 70
4.3.4 Determination of diffusion coefficients by NMR . . . . . . . . 72
4.3.5 Release of FITC-dextrans . . . . . . . . . . . . . . . . . . . . 74
4.4 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75
5 Biodegradable hydrogels for time-controlled release 77
5.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79
5.2 Materials and methods . . . . . . . . . . . . . . . . . . . . . . . . . . 81
5.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81
5.2.2 Synthesis of PEG-amines . . . . . . . . . . . . . . . . . . . . . 82
5.2.3 Synthesis of non-degradable PEG-succinimidyl carbonates . . 82
5.2.4 Synthesis of degradable PEG-succinimidyl carbonates . . . . . 83
5.2.5 Synthesis of alanine-modified PEG-amines . . . . . . . . . . . 84
5.2.6 Synthesis of 6-aminohexanoic acid-modified PEG-amines . . . 85
5.2.7 Synthesis of lysine-modified PEG-amines . . . . . . . . . . . . 86
5.2.8 Preparation and rheological characterization of hydrogels . . . 86
5.2.9 Equilibrium swelling of hydrogels and determination of network
parameters . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87
5.2.10 Degradation of hydrogels . . . . . . . . . . . . . . . . . . . . . 88
5.2.11 Mobility of incorporated macromolecules determined by fluo-
rescence recovery after photobleaching (FRAP) . . . . . . . . 89
5.2.12 Release of FITC-BSA and lysozyme . . . . . . . . . . . . . . . 90
5.2.13 Statistical analysis . . . . . . . . . . . . . . . . . . . . . . . . 90
5.3 Results and discussion . . . . . . . . . . . . . . . . . . . . . . . . . . 91
5.3.1 Physicochemical characterization of hydrogels . . . . . . . . . 91
5.3.2 Degradation of hydrogels . . . . . . . . . . . . . . . . . . . . . 94
5.3.3 Mobility of incorporated macromolecules . . . . . . . . . . . . 95
5.3.4 Release of FITC-BSA and lysozyme . . . . . . . . . . . . . . . 97
5.4 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99
6 Biointeractive hydrogels for adipose tissue engineering 101
6.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103
vii
Contents
6.2 Materials and methods . . . . . . . . . . . . . . . . . . . . . . . . . . 104
6.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104
6.2.2 Synthesis of amine-reactive polymers . . . . . . . . . . . . . . 106
6.2.3 Synthesis of collagenase-sensitive polymers . . . . . . . . . . . 106
6.2.4 Synthesis of non-degradable polymers . . . . . . . . . . . . . . 108
6.2.5 Rheological characterization of hydrogels . . . . . . . . . . . . 108
6.2.6 Equilibrium swelling of hydrogels . . . . . . . . . . . . . . . . 109
6.2.7 Degradation of hydrogels . . . . . . . . . . . . . . . . . . . . . 110
6.2.8 Cell seeding and cell culture . . . . . . . . . . . . . . . . . . . 110
6.2.9 Quantitative analysis of intracellular triglyceride accumulation 112
6.2.10 DNA assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112
6.2.11 Oil red O staining . . . . . . . . . . . . . . . . . . . . . . . . . 112
6.2.12 Statistics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113
6.3 Results and discussion . . . . . . . . . . . . . . . . . . . . . . . . . . 113
6.3.1 Physicochemical characterization of hydrogels . . . . . . . . . 113
6.3.2 Degradation of hydrogels . . . . . . . . . . . . . . . . . . . . . 116
6.3.3 Adipogenic differentiation of 3T3-L1 preadipocytes . . . . . . 119
6.4 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123
7 Summary and conclusions 125
Appendix 157
Curriculum vitae 165
List of publications 167
Acknowledgments 171
viii
Hydrogels for regenerative medicine:
Development and characterization
So eine Arbeit wird eigentlich nie fertig, man
muß sie fur fertig erklaren, wenn man nach Zeit
und Umstanden das Moglichste getan hat.
(Johann Wolfgang von Goethe)
1
Chapter 1
Introduction and goals of the thesis
3
Chapter 1 Introduction and goals of the thesis
1.1 Principles of regenerative medicine
The human body has a remarkable capacity to regenerate aged cells and damaged
tissues. After traumatic injuries and severe diseases, however, the regenerative power
of adult tissues is often not sufficient to cope with the occurred damage. Occlusion of
coronary arteries, for example, will result in necrosis of myocardial tissue and scar
formation. Depending on the size of the affected area, this may lead to heart failure
or cardiac arrest. Consequently, irreparably damaged tissues or organs have to be
replaced with artificial devices, autologous grafts, or donor organs [1–3]. However,
despite many advances in this field, medical devices (such as artificial hearts, for
example) often cannot replace the lost organ completely. The necessity of alternative
strategies is further illustrated by the ever growing mismatch between supply and
demand of organs and tissues for transplantation. In the Eurotransplant region, for
example, 7,293 people received transplants in 2008, while 15,864 people were awaiting
them [4]. The present situation will even intensify in the future, since the average
age of the Western population is increasing, and with it the incidence of age-related
diseases such as osteoporosis, diabetes, and cardiovascular diseases.
Regenerative medicine promises to overcome this dilemma. This interdisciplinary
field emerged more than two decades ago to work toward the common goal of the
repair or replacement of cells, tissues, and organs [1–3]. The original approach was
to isolate living cells from patients or other human donors, to expand them in vitro
using polymeric scaffolds, and then to re-implant the tissue-like constructs into the
patient [1]. But despite many advances, none of these cell-laden scaffolds have resulted
in complete restoration of normal tissue function [2, 3]. The complexity of growing
functional tissues in vitro has obviously been underestimated and the replacement
of whole organs is still a distant milestone in which current studies are laying the
necessary groundwork.
The currently investigated strategies are perhaps less complex, but certainly not
less ambitious. In the case of tissues with inherent regenerative capacities, tissue
regeneration can be promoted by inserting an appropriate biomaterial to prevent
undesirable, rapidly proliferating cells from entering the site of defect. In another
approach, research tries to stimulate the body’s own repair mechanisms by mimicking
the regulatory function of growth factors. This will require the development of “smart”
4
1.2 Biomaterials for regenerative medicine
biomaterials that allow delivering growth factors in a spatio-temporally controlled
manner [5–7]. Recently, nanostructured biomaterials have also been proposed that
target to the injury site and self-assemble into higher order scaffold structures. These
would provide an appropriate microenvironment to recruit and activate endogenous
stem cells to form differentiated tissues and organ structures [3].
1.2 Biomaterials for regenerative medicine
Biomaterials play a central role in regenerative medicine and tissue engineering as
carrier systems for drug molecules or cells. Commonly used biomaterials include
biodegradable poly(glycolic acid) (PGA), poly(lactic acid) (PLA), and poly(lactide-
co-glycolide) (PLGA). However, these polymers are rather hydrophobic and are
typically processed under relatively harsh conditions, which makes the incorporation
of fragile biomolecules or living cells a challenge. Most of the fabricated scaffolds were
designed to withstand mechanical loads and to degrade within an appropriate period
of time, while additional functionalities (such as the ability to trigger specific cellular
responses) have often been neglected. During the past decade, however, there has
been a substantial paradigm shift in the design criteria of modern biomaterials [8].
Current developments integrate principles from cell and molecular biology to mimic
certain aspects of the natural extracellular matrix (ECM).
Therefore, hydrogels have been proposed as potential alternatives for a variety of
drug delivery and tissue engineering applications [7, 9–13]. These hydrophilic polymer
networks absorb large amounts of water and demonstrate excellent biocompatibility
due to their physicochemical similarity to the native ECM. Furthermore, gel for-
mation usually proceeds at ambient temperature without requiring organic solvents.
Hydrogels can be classified into natural, synthetic, and composite gels according to
their origin and composition. Furthermore, one can distinguish between ‘reversible’
or ‘physical’ gels and ‘permanent’ or ‘chemical’ gels. In physical gels, networks are
held together by molecular entanglements and/or secondary forces including ionic
interactions, hydrogen bonds, and hydrophobic interactions. In contrast, chemical
gels are characterized by covalent cross-links between the individual polymer chains.
5
Chapter 1 Introduction and goals of the thesis
In terms of industrial production processes, hydrogels derived from synthetic
polymers are especially appealing, as their chemical and physical properties are easily
controlled and reproduced. Synthetic materials include poly(vinyl alcohol) (PVA),
derivatives of poly(acrylic acid) (PAA), poly(ethylene glycol) (PEG), and synthetic
polypeptides. Of these substances, polymers derived from PEG are among the most
commonly applied hydrogel-forming materials. The widespread use of these polymers
primarily results from their excellent biocompatibility and high solubility in water and
organic solvents [14]. The versatility of the PEG macromer chemistry further allows
for the design of ‘biomimetic’ hydrogels that mimic the complexity of the natural
ECM [10, 11, 13]. These hydrogels can be equipped with molecular cues that guide
the adhesion and/or recruitment of cells (e.g. endogenous stem cells), degradation
sites for cellular proteases to allow for cell-triggered remodeling, and binding sites for
soluble signaling molecules (such as growth factors or cytokines). Altogether, PEG is
deemed to be an ideal starting material for the development of sophisticated hydrogel
systems for drug delivery and regenerative medicine applications.
1.3 Goals of the thesis
This thesis is focused on the development and characterization of PEG-based hydrogels
for controlling drug delivery and promoting tissue regeneration. To achieve these goals,
cross-linking methods had to be identified that are sufficiently gentle to be performed
in the presence of cells or in vivo. This would facilitate injection and provide an
effective way to encapsulate drug molecules or living cells (Figure 1.1). Based on the
established chemistry, different polymers were synthesized as building blocks for the
preparation of hydrogels. As a consequence of this combinatorial approach, a variety
of hydrogels could be prepared from comparatively few macromers. The developed
hydrogels may serve as inert space-filling agents, as carrier systems for the controlled
release of drug molecules, or as three-dimensional scaffolds in cell-based approaches.
Similar to the native ECM, synthetic biomaterials must provide an “instructive”
microenvironment that directs cell proliferation and differentiation [8, 15–17]. For
6
1.3 Goals of the thesis
these reasons, an exact knowledge of the chemical, physical, and topographical factors
guiding tissue morphogenesis in vitro and in vivo will be crucial.
OO O
O
N
O
O
NH2
OO
OO N
H
O
O
O
Component A(amine-reactive)
Component B(amine-containing)
Hydrogel(covalently cross-linked)
Figure 1.1: Principle of in situ forming hydrogels. An amine-reactive compound (A) iscombined with an amine-containing component (B) e.g. by using a two-chamber syringe.Directly after mixing the liquid precursor solutions, the individual polymer chains arecross-linked to form a highly elastic hydrogel. Cells or drug molecules can be easilyincorporated by suspending or dissolving them in one of the two precursor solutions.
In Chapter 2, the influence of environmental cues on cell proliferation and differ-
entiation is, therefore, reviewed. Since most PEG-based hydrogels are biologically
and chemically inert [10], the impact of physical factors on cell behavior is primarily
stressed. To this end, the physical properties of hydrogels will be discussed, which
include their gel forming characteristics, their mechanical properties, and degradation
behavior. A short introduction to methods of characterizing the mechanical properties
of hydrogels is also included. The chapter is completed by a detailed review of several
in vitro studies that illustrate the complex interplay between substrate stiffness,
degradability, cell differentiation, and tissue morphogenesis.
According to these theoretical considerations, the following work was focused on the
development of in situ forming hydrogels for intraocular applications (Chapter 3).
Age-related macular degeneration (AMD) and proliferative diabetic retinopathy (PDR)
are among the leading causes of blindness in industrialized nations [18]. PDR is
characterized by an abnormal growth of blood vessels into the vitreous body, a
7
Chapter 1 Introduction and goals of the thesis
virtually acellular, gel-like network of collagen fibrils and glycosaminoglycans that
fills the posterior segment of the eye. In these patients, a total replacement of
the affected vitreous may be required in order to prevent blindness or to restore
vision. However, today’s clinically used substitutes differ significantly from the natural
vitreous body with regard to their physicochemical properties and mechanics. Some
of these substances are also associated with severe side-effects when kept intravitreally
over longer periods of time. Therefore, this work was aimed at developing a better
tolerated, hydrogel-based vitreous substitute with mechanical properties similar to
those of the natural vitreous body. Particular attention had to be paid to the
biocompatibility, optical transparency, and injectability of the proposed hydrogels.
The developed hydrogels are non-degradable and designed to act as inert space-filling
agents over longer periods of time.
Apart from their potential application as vitreous substitutes, hydrogels would also
be promising materials for the delivery of drugs to the posterior segment of the eye (e.g.
to prevent the above described neovascularization). However, despite many favorable
characteristics, hydrogel-based drug delivery systems still have some limitations. In
fact, the high water content of most hydrogels often results in relatively rapid drug
release over several hours to a few days. To overcome these limitations, efforts were
made to extend the duration of drug release and to expand the range of molecules
which can be effectively delivered by hydrogels (e.g. by increasing the average network
mesh size). Since the resulting release profiles are hardly predictable, newly developed
drug delivery systems are usually characterized by release experiments. However, these
experiments are time-consuming and their reliability is often limited. In the next study,
the significance of mechanical testing, swelling studies, fluorescence recovery after
photobleaching (FRAP), and pulsed field gradient nuclear magnetic resonance (NMR)
spectroscopy was, therefore, investigated for the characterization of hydrogel-based
drug delivery systems (Chapter 4).
To prolong the release of incorporated peptides or proteins, the existing hydrogels
had to be modified. For this purpose, the possibility of tethering drug substances to the
hydrogel backbone was investigated. Ideally, hydrogel cross-linking and drug tethering
would be performed simultaneously without requiring chemical modifications of the
drug molecules. This would improve handling and flexibility of the developed drug
delivery system, since any peptide or protein could be incorporated by simply dissolving
8
1.3 Goals of the thesis
them together with the gel-forming polymers. To achieve this goal, biodegradable
polymers were synthesized that readily react with amino groups of other polymers,
peptides, or proteins (Chapter 5). During cross-linking, the drug molecules are
covalently bound to the gel network, which effectively prevents their immediate release.
Release kinetics is then controlled by the degradation of the anchor group; drug
diffusivity only plays a secondary role. The anchor groups used for drug conjugation
had to be carefully designed in order to prevent potential loss of bioactivity and
to allow for the time-controlled release of incorporated molecules. The developed
hydrogels were characterized by mechanical testing, the established FRAP technique,
and release experiments.
Besides their use as inert space filling agents (Chapter 3) and drug delivery
systems (Chapter 4 and 5), in situ forming hydrogels could also be applied as
three-dimensional scaffolds in cell-based approaches. The developed hydrogels provide
for effective cell encapsulation and unrestricted diffusion of nutrients and metabolites.
For a successful application in regenerative medicine, hydrogel scaffolds must bear
the occurring mechanical loads and provide a suitable microenvironment to promote
cell proliferation and differentiation. Once placed at the application site, the scaffold
should degrade in spatial and temporal synchrony with the formation of new tissue.
For this purpose, the gel-forming polymers were functionalized with a synthetic
tetrapeptide (Ala–Pro–Gly↓Leu) to make them susceptible to proteolytic breakdown
(Chapter 6). These cell-responsive hydrogels mimic the proteolytic recognition of
the natural ECM and are degraded by cell-secreted proteases. In the last study, these
biointeractive hydrogels were seeded with 3T3-L1 preadipocytes to investigate the
impact of substrate stiffness, adhesiveness, and degradability on cell proliferation and
differentiation.
9
Chapter 2
Rational design of hydrogels fortissue engineering: Impact of physicalfactors on cell behavior
Ferdinand Brandl1, Florian Sommer1,2, Achim Gopferich1
1 Department of Pharmaceutical Technology, University of Regensburg, 93040 Regensburg2 Boehringer Ingelheim Pharma GmbH & Co. KG, 88397 Biberach an der Riß
Published in Biomaterials 28 (2), 134–146 (2007).
11
Chapter 2 Rational design of hydrogels for tissue engineering
Abstract
When designing suitable biomaterials for tissue engineering applications, biological
and chemical parameters are frequently taken into account, while the equally impor-
tant physical design variables have often been neglected. For a rational design of
biomaterials, however, all variables influencing cell function and tissue morphogen-
esis have to be considered. This review will stress the development of cross-linked
hydrogels and outline the impact of their physical properties on cell function and
tissue morphogenesis. In the first part, the principles of cellular mechanosensitivity,
as well as the influence of substrate mechanics on cell behavior, will be discussed.
Afterwards, methods to characterize the mechanical properties of biomaterials will be
presented. The subsequent chapters will address hydrogels that allow for the control
of their physical qualities followed by a discussion of their use in tissue engineering
applications.
12
2.1 Introduction
2.1 Introduction
The human organism is composed of around 1013 cells that are classified into more
than 200 different cell types [19]. Cell function, tissue morphogenesis, and organ
development are thought to be regulated by a fine-tuned interplay of chemical, physi-
cal, and topographical factors [19–21]. Many of the principles guiding embryogenesis
in vivo are also considered to be involved in the regulation of tissue development
in vitro. Despite the proliferation of this concept, the design of biomaterials for
tissue engineering is still frequently guided by the principles of trial and error, rather
than by rational considerations of the specific demands. Many biomaterials have
been developed to meet particular biological and chemical requirements (e.g. biocom-
patibility, degradability, mediation of cell adhesion, etc.). Other design parameters,
such as the physical properties of the biomaterial, were regarded with respect to the
processing conditions, the mechanical load capacity, or the diffusivity of solutes, but
not with respect to the biological response. This is probably at least in part due
to the lack of adequate methods of measuring the physical attributes of tissues or
tissue-engineered constructs.
For a rational design of biomaterials, however, all variables influencing cell function
and tissue morphogenesis have to be considered. To understand the influence of
each parameter, their individual signaling pathways have to be elucidated. Together,
these fundamentals will reveal “set screws” for the design of biomaterials. Adjusting
these parameters to the requirements of each specific application would allow for the
creation of “custom-made” biomaterials that direct the development of desired tissues.
Thereby, the inherent characteristics of biological tissues may serve as guides for this
process [19–24].
This review was written to promote the rational design of hydrogels for tissue
engineering applications with a special emphasis on physical properties. Hydrogels are
highly hydrated networks that have been fabricated from a wide range of hydrophilic
polymers [9, 10, 25]. They can be classified into ‘reversible’ or ‘physical’ gels and
‘permanent’ or ‘chemical’ gels. In physical gels, networks are held together by
molecular entanglements and/or secondary forces including ionic cross-links, hydrogen
bonds, and hydrophobic interactions. In contrast, chemical gels consist of covalently
cross-linked networks [9].
13
Chapter 2 Rational design of hydrogels for tissue engineering
In the first part of this review, we will discuss the basic principles of cellular mechano-
sensitivity. Theoretical considerations are illustrated by in vitro studies that elucidate
the general cell responses on two-dimensional model substrates. Subsequently, we
will outline the problems of characterizing the mechanical properties of biological
tissues and hydrogels followed by a discussion on the rational design of hydrogels
for tissue engineering applications. Finally, we will stress the impact of mechanical
characteristics and degradability on cell function and tissue morphogenesis. For this
purpose, we will present relevant in vitro studies as well as available in vivo data.
2.2 Environmental factors as morphogenetic guides
In tissues, cells are embedded within the extracellular microenvironment, a highly
hydrated network that comprises three classes of stimuli or cues that stem from
the following sources: insoluble hydrated macromolecules (e.g. fibrillar proteins,
proteoglycans, or polymer chains), soluble molecules (e.g. growth factors or cytokines),
and membrane-associated molecules of neighboring cells [8, 24]. As it is assumed that
most interactions between cells and these extracellular effectors are determined by
associations between receptors and corresponding ligands [26], we will concentrate
here upon specific ligand-receptor interactions and disregard nonspecific effects, such
as electrostatic interactions. Ligand-receptor interactions are considered as specific, as
they depend on detailed topographical features of interacting structures (“lock-and-key
principle”) [26].
Soluble receptor ligands, such as growth factors and cytokines, are thought to diffuse
to their target receptors. The transmitted information will arise from the type of
signaling molecule as well as its local concentration [19]. The resulting cellular response
to that kind of stimulus is currently being investigated in detail; comprehensive reviews
dealing with the application of growth factors in tissue engineering can be found
in the literature [27–29]. By contrast, the pure biochemical information provided
by ligands attached to an extracellular structure, such as the extracellular matrix
(ECM), is supplemented by additional degrees of information including the spatial
distribution of ligands and the mechanical properties of the structure the ligands
are attached to [19]. Spatial variations in adhesiveness, for example, can lead to
a directed cell movement towards regions of higher ligand density, a phenomenon
14
2.2 Environmental factors as morphogenetic guides
termed haptotaxis [30–32]. In the following paragraphs, however, we will focus on
the impact of mechanical cues on such cell behavior.
2.2.1 Integrins as mechanoreceptors
In the past, great efforts have been made to elucidate how physical forces, applied
to either the ECM or the cell surface, induce biochemical alterations inside the cell.
Today, there is much evidence that mechanical signals are transferred into the cell
across transmembrane molecules, such as integrins, which couple extracellular anchors
to the cytoskeleton [33–35]. Integrins constitute a large family of transmembrane,
heterodimeric receptors that bind to specific amino acid sequences, such as the
arginine–glycine–aspartic acid (RGD) recognition motif, present in all major ECM
proteins [36]. After binding to ECM ligands, integrins cluster together to form dot-
like adhesive structures termed focal complexes. Depending on the stiffness of the
underlying substrate, focal complexes can disappear or evolve into focal adhesions.
These multi-molecular plaques anchor bundles of actin filaments (stress fibers) and
mediate strong adhesion to the substrate. In turn, focal adhesions are considered
to be a source for fibrillar adhesions, which are involved in matrix assembly into
extracellular fibrils [37, 38]. Studying cell-matrix interactions in a three-dimensional
(3-D) context, Cukierman et al. described distinctive “3-D matrix adhesions” that
differed from both focal and fibrillar adhesions characterized on two-dimensional
(2-D) substrates in structure, localization, and function. They further speculated
that classically described in vitro adhesions are exaggerated precursors of those, more
biologically relevant “3-D matrix adhesions” [39, 40].
To explain the molecular basis of mechanotransduction, Ingber et al. proposed the
cellular tensegrity model [34, 35]. According to this model, living cells are thought
to exist in a state of pre-stress, actively generated by myosin-II driven isometric
contractions of the actin cytoskeleton. Structural elements that resist compression,
notably internal microtubule struts and ECM adhesions, act as a counterbalance. As
cell-ECM adhesions and microtubule struts resist cytoskeletal tension in a comple-
mentary manner, changes in ECM mechanics or extracellular perturbations generate
mechanical forces within the cytoskeletal structure. In reaction to unbalanced forces,
cells rearrange cell-matrix adhesions, reorganize their cytoskeleton, and immediately
shift their shape. For example, if the stiffness of the ECM exceeds the stiffness of the
15
Chapter 2 Rational design of hydrogels for tissue engineering
cytoskeleton, cells will flatten and spread. In the opposite case, cells will retract or
become rounded. Biochemical responses are thought to be mediated by conforma-
tional changes of regulatory molecules within the adhesion plaque. These molecular
events, in turn, trigger signal transduction cascades which ultimately regulate cell
proliferation, differentiation, and apoptosis [33, 35].
2.2.2 Mechanical cues regulate cell behavior
Model considerations of cellular mechanosensitivity are also supported by experi-
mental data. Using fibronectin-coated beads held in an optical trap, Choquet et al.
demonstrated that cells strengthen their integrin-mediated contacts to the beads in
proportion to the force restraining it. According to the authors, this mechanism might
allow cells to migrate through the ECM in response to its mechanical properties [41].
In a later study, Lo et al. verified the idea of mechanotaxis by demonstrating that cell
movement is guided by the rigidity of the substrate [42]. Similar results were found
by Gray et al. They used a micropatterning technique to produce fibronectin-coated
surfaces of varying stiffness and observed cell migration towards stiffer regions of the
substrate (Figure 2.1) [43].
Figure 2.1: NIH/3T3 fibroblasts cultured on fibronectin-coated poly(dimethylsiloxane)(PDMS) substrates. The squares are stiff, whereas the regions surrounding the squaresare compliant. Accumulation of cells on stiffer regions was found to be due to migration,not proliferation, of cells in response to the mechanical patterning (mechanotaxis). Scalebars represent 100 µm. Reprinted with permission from Gray et al. [43]. c© 2003 JohnWiley & Sons, Inc.
In addition to migration, a variety of other cell functions, such as cell spreading,
growth, and differentiation, are also modulated by the substrate mechanics. Pelham et
16
2.3 Mechanical properties of materials and their characterization
al. reported that cells on flexible substrates showed reduced spreading and increased
rates of motility compared to cells on rigid substrates [44]. Wang et al. found cell
proliferation to be increased on culture substrates of higher mechanical stiffness. In
contrast, the rate of apoptosis was increased on more flexible substrates [45]. Studying
angiogenesis in vitro, Vailhe et al. demonstrated that the formation of capillary-like
structures was influenced by the rigidity of the fibrin gels utilized [46]. Similar
results were obtained be Deroanne et al., who could show that cell differentiation
was affected by the mechanical properties of the supportive matrix: with decreasing
substrate rigidity, the number of endothelial cells switching to a tube-like pattern
increased [47]. Differentiation of neuronal cells also seems to be regulated by the
mechanical properties of the culture substrate. According to Flanagan et al., the
formation of neurite branches was enhanced by softer substrates [48].
As mechanosensitivity is related to cells’ ability to rearrange adhesion ligands
presented by the substrate and to apply traction forces to the material [49], substrate
mechanics and adhesiveness should be regarded as coupled variables. Rowley et al.
reported that myoblast differentiation on alginate gels was regulated by the mechanical
properties of the substrate as well as the RGD density [50]. Investigating spreading
of smooth muscle cells (SMCs) on collagen-coated polyacrylamide gels, Engler et
al. showed matrix compliance and ligand density to be highly coupled variables
that determine mean cell responses [51]. Finally, Peyton and Putnam reported a
biphasic dependence of cell migration speed on ECM stiffness. In their study, the
optimal stiffness at which cell migration speed is maximized was found to depend on
the density of immobilized ECM ligands [52]. For more detailed information about
the crucial role of substrate mechanics and adhesiveness in cell regulation, several
comprehensive reviews are recommended [19–21].
2.3 Mechanical properties of materials and theircharacterization
The biochemical (e.g. adhesiveness) and physical properties (e.g. substrate stiffness)
of the extracellular microenvironment have been recognized as interdependent fac-
17
Chapter 2 Rational design of hydrogels for tissue engineering
tors that influence cell function and tissue morphogenesis in multiple ways [19–21].
Consequently, both biochemical and physical characteristics must be considered when
designing hydrogels for tissue engineering applications [10, 25]. Hydrogels will act
as morphogenetic guides if their biochemical and physical attributes are tailored
to provide an appropriate environment for cell adhesion, migration, growth, and
differentiation [8, 15–17, 24]. To determine the optimal parameters, the mechanical
properties of tissues or remodeled ECM may serve as reference points [20, 21]. This,
in turn, will require accurate methods of measuring the mechanical properties of
tissues, fabricated hydrogels, and tissue-engineered constructs.
Amongst other methods, the mechanical properties of materials, including tis-
sues [53] and hydrogels [54], are characterized by tensile tests, compression tests, and
dynamic mechanical analysis (DMA). For uniaxial tensile testing, dog bone-shaped
samples are placed between two clamps and stretched at constant extension rates.
From these experiments, the Young’s modulus of the material can be determined.
It is defined as the ratio of tensile stress to tensile strain, whereas the maximal
tensile stress carried by a material is defined as the tensile strength. Similarly, the
compressive modulus is defined as the ratio of compressive stress to compressive strain.
Testing is performed by uniaxial compression of cylindrical specimens between two
smooth impermeable platens (unconfined compressive testing). In contrast to that,
confined compressive testing is carried out in a confining chamber where the sample
is loaded by a permeable piston. These experiments reveal the aggregate modulus of
the material. Depending on the applied testing mode, the calculated values of the
Young’s modulus will differ: frictional effects and/or interdigitation of the sample into
the platen pores may increase the moduli obtained in confined compression [55]. The
compressive strength is defined as the maximal compressive stress that a sample can
withstand. Both, Young’s modulus and compressive modulus are a measure of the
stiffness of a given material, which mirrors the resistance of an elastic body against
the deflection of an applied force.
DMA is typically performed to measure the viscoelastic behavior of materials. In
rheological terms, ‘viscoelastic’ means the concomitance of viscous (“liquid-like”)
and elastic (“solid-like”) behavior. For a given material, the proportion of viscous
to elastic properties will depend on the experimental conditions (e.g. timescale and
temperature). DMA assessments require the application of a sinusoidal shear load on
18
2.3 Mechanical properties of materials and their characterization
the sample. A stress transducer measures the applied shear stress (σ∗). The strain
induced in the sample (γ∗) is measured using a strain transducer. The complex shear
modulus G∗ is defined as follows:
G∗ = G′ + i ·G′′ = σ∗
γ∗(2.1)
G′ is referred to as the real part of G∗ (also elastic or storage modulus) and represents
the relative degree of a material to recover (“elastic response”). G′′ is referred to as
the imaginary part of G∗ (also viscous or loss modulus) and represents the relative
degree of a material to flow (“viscous response”) [56]. Measuring G∗ against the shear
stress or shear strain, respectively, allows to determine the stiffness and strength of a
given material.
Using tensile tests, compressive tests, or DMA, the elastic moduli of various
tissues have been determined (Table 2.1) [53]. In general, the measured moduli
range over several orders of magnitude; neuronal tissue [57] is much softer than
cartilaginous tissue [58] or bone tissue [59], for example. However, the obtained
values should be regarded just as rough estimates for the mechanical characteristics of
biological tissues. Nevertheless, the observed differences imply that distinct mechanical
microenvironments exist for different cell types and tissues [21].
Table 2.1: Mechanical properties of different biological tissues. Many other studies can befound elsewhere [53].
Specimen Testing method Results Ref.
Bovine spinal cord(gray matter)
Tensile test Tangent modulia,b ranged between 63.9± 7.9and 112.3± 10.2 kPa depending on the strainrate
[57]
Articular cartilagefrom human hip joints(femoral head)
Biphasic creepindentation test
Aggregate modulia ranged between0.679± 0.162 and 1.816± 0.868 MPadepending on the location
[58]
Cortical bone fromhuman femoraldiaphysis
Tensile test Total average value of the Young’s modulus:17.9 GPa (data obtained at strain rates of4 · 10−2 s−1)
[59]
aData are given as mean ± standard deviation.bThe tangent modulus is defined as the slope of the tangent to the stress-strain curve at a specific
point. Within the linear elastic region, the tangent modulus is equal to the Young’s modulus.
19
Chapter 2 Rational design of hydrogels for tissue engineering
In biological tissues local regions of high stiffness exist beside regions that exhibit
much lower values for the elastic modulus. These heterogeneities are due to the
composite character and the ongoing remodeling of the ECM [20]. Admittedly, local
differences in the mechanical properties will not be detected by bulk measurements,
such as tensile tests, compressive tests, and DMA. But because cells respond to spatial
variations in the substrate stiffness, which can be on the order of microns [19, 20, 43],
the local mechanical properties rather than the bulk properties will be crucial for
the design of hydrogels. In addition, tensile tests, compressive tests, and DMA may
affect the structural integrity of the sample or even involve its destruction [54]. When
surveying the mechanical properties of living tissues or tissue-engineered constructs,
however, non-invasive and non-destructive methods with high spatial resolution would
be preferred. In the next paragraphs, we will highlight some of these methods.
2.3.1 Atomic force microscopy
Atomic force microscopy (AFM) can be used not only for imaging the topography
of surfaces, but also for measuring forces on a molecular level. To investigate the
mechanical properties of soft matrices or thin films, the sample is compressed by the
indenting AFM tip (Figure 2.2). The loading force is calculated from the deflection
and the spring constant of the cantilever. To calculate the Young’s modulus of the
material, force-indentation-curves are recorded and fitted to the Hertz model, which
describes the elastic deformation of two spherical surfaces under load [60, 61].
Engler et al. used AFM to investigate the mechanical environment seen by SMCs
in vivo and correlated this with SMC responses on collagen-coated polyacrylamide
(PAAM) gels. Surface probe measurements within the SMC-rich medial layer of
sectioned arteries revealed an apparent Young’s modulus of ∼ 5 – 8 kPa; the Young’s
moduli of collagen-coated PAAM gels ranged between ∼ 1 kPa and ∼ 35 kPa. Spread-
ing of SMCs on PAAM gels showed a hyperbolic dependence on the elastic modulus
of the substrate. Remarkably, half-max spreading of SMCs occurred on gels that
approximated the stiffness of the arterial media (E 12−spread ≈ Emedia). For this reason,
E 12−spread is regarded as a mechanical set point for SMCs. Engler et al. concluded from
20
2.3 Mechanical properties of materials and their characterization
x
y z
Lase
r
PSPD
Cantilever with tip
Piezo scanner
Sample surface
Feedback electronic
Figure 2.2: Diagram of AFM instrumentation [61]. A sharp tip at the free end of amicroscale cantilever is used to probe the sample surface. The sample is mounted on apiezoelectric scanner that moves the sample in the x and y directions for scanning thesurface and in the z direction for indenting the sample. A laser beam reflected from theback of the cantilever onto a position sensitive photodiode (PSPD) forms an optical leversystem that measures the deflection of the cantilever. From this data and the springconstant of the cantilever the loading force can be calculated.
these experiments that surface probe measurements allow for an accurate assessment
of the local mechanical properties of various materials including biological tissues [62].
2.3.2 Magnetic resonance elastography
Magnetic resonance elastography (MRE) is a non-invasive and non-destructive tech-
nique that visualizes spatial changes in mechanical properties. It has been successfully
used to characterize the elastic properties of gel samples and tissue explants ex vivo.
But MRE also provides information about the mechanical properties of soft tissue in
vivo, which allows for the detection of pathological changes, such as soft tissue tumors,
by a sensitive and safe method [63–65]. In this method, shear waves are generated
within the sample using an electromechanical actuator coupled to the surface of
the object. Using a magnetic resonance imaging (MRI) system with an additional
motion sensitizing gradient, the displacement patterns corresponding to the shear
21
Chapter 2 Rational design of hydrogels for tissue engineering
waves can be measured (Figure 2.3). The obtained “wave images” directly visualize
the propagation of shear waves within the sample and allow the reconstruction of
viscoelastic parameters at each location in the material [63, 64].
Gvib
RF
Gslice
Gphase
Gread
Ima
gin
gg
rad
ien
ts
Trigger pulses
Oscillator/Amplifier
Actuator coil
Pivot
Motiondirection
Direction of motion-sensitizing gradient
Sample
MRI system (shaded) with additionalmotion-sensitizing gradient (G )vib
Figure 2.3: Schematic diagram of the MRE system [63]. A conventional MRI systemoperating with imaging gradients (Gslice, Gphase, and Gread) and radiofrequency (RF)pulses is equipped with an additional motion-sensitizing gradient (Gvib) (left). Theimaging gradients are used to encode the spatial positions of the MR signal. Triggerpulses provided by the imager synchronize an oscillator that drives an electromechanicalactuator coupled to the surface of the sample (right). In the presence of Gvib, the cyclicmotion of the spins causes a measurable phase shift in the received MR signal. From thisphase shift, the displacement in each volume element can be calculated. The data thusobtained are used to visualize the propagating shear waves within in the sample and toreconstruct the corresponding viscoelastic parameters.
Clinical magnetic resonance (MR) systems typically provide a spatial resolution of
1 mm× 1 mm× 10 mm, which would not be appropriate to survey the mechanical
properties of small tissue-engineered constructs. In a recently published work, however,
Othman et al. reported the development of an enhanced MRE method termed micro-
scopic magnetic resonance elastography (µMRE). This technique has been used to im-
age shear wave propagation with a microscopic resolution of 34 µm×34 µm×500 µm.
To evaluate the potential of µMRE for identifying the mechanical properties of
22
2.3 Mechanical properties of materials and their characterization
tissue-engineered constructs, Othman et al. cultured human bone marrow stromal
cells (BMSCs) on gelatin sponges and differentiated them either into adipogenic
or osteogenic cells. In preliminary experiments using µMRE, the shear stiffness of
adipogenic and osteogenic constructs was estimated to be ∼ 1.2 and ∼ 15 kPa, respec-
tively. Although the algorithms used to reconstruct the material’s properties still had
to be adapted, the µMRE technique provides a valuable tool to monitor the mechanical
properties of tissue-engineered constructs during growth and differentiation [66].
2.3.3 Monitoring of cellular traction forces using fluorescenceresonance energy transfer
Kong et al. [67] proposed a fluorescence resonance energy transfer (FRET) technique
that may be adapted to study cell-material mechanics in three-dimensional culture.
FRET occurs between a donor fluorochrome and an acceptor fluorochrome, if the
emission wavelength of the donor and the excitation wavelength of the acceptor
overlap. Furthermore, the spatial distance between donor and acceptor has to be less
than 10 nm, such that the former can transfer energy to the latter (Figure 2.4) [68–70].
In their study, Kong et al. [67] coupled RGD-containing oligopeptides to sodium
alginate and labeled the immobilized peptides with either Alexa Fluor 488 (green
fluorescence) or Alexa Fluor 546 (red fluorescence). Hydrogels prepared by cross-
linking equal volumes of differently labeled polymers with calcium were seeded with
murine preosteoblasts and incubated in medium. Imaging was performed by laser
scanning microscopy (excitation wavelength 488 nm). In these experiments, red
fluorescence was limited to regions containing adherent cells, indicating that the
labeled peptides not involved in cell adhesion were separated by a greater spacing
than the critical distance required for FRET (Figure 2.4). With increasing substrate
stiffness, the yield of red fluorescence first increased and then decreased. This is
related to the capability of cells to cluster the adhesion peptides. The calculated force
that cells exerted to displace the adhesion peptides, however, increased in proportion
to the substrate stiffness. These results correlate very well with observed changes in
cell phenotype, which have been reported to depend on cell adhesion stiffness. The
FRET technique is, therefore, regarded as a molecular ruler to monitor displacements
23
Chapter 2 Rational design of hydrogels for tissue engineering
488 546 546488FRET
lex = 488 nm
lem = 580 – 620 nm
lex = 488 nm
A B
Critical distancerequired for FRET
lem = 500 – 540 nm
No FRET
Figure 2.4: FRET is a process in which energy is transferred nonradiatively from an exciteddonor fluorophore to an acceptor fluorophore. It occurs with measurable efficiency if thetwo fluorophores are situated less than 10 nm apart [68–70]. (A) Excitation of the sample(λex = 488 nm) results in green fluorescence (λem = 500− 540 nm), as the correspondingfluorophores (RGD-containing oligopeptides labeled with either Alexa Fluor 488 or AlexaFluor 546) are separated by a greater spacing than the critical distance required forFRET. (Alexa Fluor 546 is not excited at this wavelength.) (B) Seeded cells rearrangeadhesion molecules presented from the substrate. Excitation of the sample leads to areduction in the yield of green fluorescence, but increases the yield of red fluorescenece(λem = 580− 620 nm) [67].
between adhesion ligands and provides a valuable method to calculate cell traction
forces without mechanical or chemical manipulations.
2.4 Rational design of hydrogels for tissue engineeringconsidering physical aspects
Biochemical and physical parameters were identified as essential design variables
of hydrogels used in tissue engineering applications [8, 10, 15–17, 22–25]. In this
chapter, we will stress the physical properties of hydrogels, which include their gel
forming characteristics, their mechanical or viscoelastic properties, respectively, and
their degradation behavior. Below, we will present examples of current methods of
controlling the physical properties of hydrogels. Alginates and poly(ethylene glycol)
(PEG) serve as models, as their properties reflect those of many other gel forming
polymers as well. The following considerations, however, can be applied to other
polymers, too.
24
2.4 Rational design of hydrogels for tissue engineering
In general, all hydrogels used in biomedical applications must be biocompatible.
Because the apparent mesh size of polymeric gels is typically much smaller than a
cell’s diameter, it would be useful to introduce cells into the liquid precursors of the
gel, rather than to the preformed hydrogel itself. To accomplish this, gel forming
methods have to be chosen that can be conducted in the presence of cells or in vivo
without causing damage [10, 16, 25].
Alginates are naturally occurring polysaccharides and consist of guluronic acid (G)
and mannuronic acid (M) organized into blocks of varying composition (G-blocks,
M-blocks, and MG-blocks). Gels are formed when divalent cations (e.g. Ca 2+) in-
teract with G-blocks to form ionic bridges between different polymer chains [71].
Because of their recognized biocompatibility and gentle gelling properties, hydrogels
prepared from alginates are very attractive for many tissue engineering applica-
tions [10, 25]. PEG represents another type of polymer that is widely used in
biomedical applications [10, 25]. Aqueous solutions of PEG macromers terminated
with acrylate or methacylate groups can be photo-polymerized in the presence of
cells using UV or visible light, respectively, in combination with a proper initiating
system to form covalently cross-linked hydrogels [72]. Besides ionic interactions
and photo-polymerization, cross-linking is also accomplished by chemical reaction of
complementary groups [73]. Vinylsulfone-functionalized PEG macromers can be cross-
linked utilizing a Michael-type addition reaction between the vinylsulfone end groups
and thiol-bearing compounds (Figure 2.5). These reactions can be conducted under
physiological conditions and allow for the preparation of hydrogels in the presence of
cells or in vivo [74]. Moreover, a variety of temperature-sensitive hydrogel systems
are described in the literature [75]. Recently, important progress has also been made
to form nanofibrillar matrices in situ by molecular self-assembly of synthetic peptides
or proteins [76].
Due to their hydrophilic nature, most synthetic hydrogels are known to prevent
the adsorption of ECM proteins. In addition, non-adhesiveness is accomplished
because cells lack adhesion receptors for most hydrogel forming polymers [10, 25].
In order to design hydrogels that mediate attachment of cells, entire ECM proteins
or synthetic peptide sequences capable of binding to cellular receptors have been
covalently coupled to the polymer chains [8, 10, 15–17, 24, 25]. Incorporation of
25
Chapter 2 Rational design of hydrogels for tissue engineering
SH
NH2
Protease substrate
O
NH
OH
O
SH
OS
O
O
O SO
O
O
O
+ +
OO
S
O
OS
NH2
O
Protease substrateNH
OHO
SS
OO
O
O
Figure 2.5: Michael-type addition reaction between vinylsulfone-functionalized PEG macro-mers and cysteine containing peptides. Cross-linking with enzymatically cleavable se-quences renders the gels susceptible to proteolytic breakdown.
biologically active substances is another strategy by which hydrogels can be modified
to regulate cell function and tissue morphogenesis [8–10, 15–17].
Once placed at the application site, the hydrogel scaffolds should be able to bear
the local mechanical loads until the cells have produced their own functional ECM.
Moreover, the hydrogel should provide an appropriate mechanical environment to
support cell migration, proliferation, and differentiation [16, 74]. As each tissue
provides its own mechanical microenvironment, the mechanical characteristics of
hydrogels used in tissue engineering have to be adapted to the intended application:
engineering neuronal tissue will require other mechanical conditions than cartilage or
bone, for example. In part, the mechanical properties of hydrogels are predetermined
by the inherent characteristics of the building blocks including their chemistry and
molecular weight (MW). The gel strength can be further tailored by varying the
concentration and composition of building blocks, by altering the method of cross-
linking, and by adjusting the cross-link density or mesh size [54].
As only the G-blocks participate in ionic cross-linking, the gel strength of alginates
depends on the monomeric ratio (M:G ratio) and the length of G-blocks [71]. Further-
more, the mechanical properties and swelling degree can be regulated by controlling
the cross-link density (e.g. by altering the concentration of divalent cations) and
using different principles of cross-linking (e.g. covalent cross-linking) [77]. Increasing
the concentration of alginate also enhances the strength of alginate hydrogels [78].
26
2.4 Rational design of hydrogels for tissue engineering
Similarly, the mechanical properties of PEG gels are altered when the weight fraction
of PEG diacrylate [79] or PEG dimethacrylate [80] increases. This is explained by the
cyclization of macromers, which predominantly occurs at high solvent concentrations,
finally leading to more loosely cross-linked hydrogels [80]. Furthermore, the molecular
weight between cross-links and mesh size are also influenced by the molecular weight
of the PEG macromer [81]. In contrast to PEG diacrylates and dimethacrylates,
vinylsulfone-functionalized PEG macromers are typically branched. The mechanical
properties and swelling ratio of hydrogels formed by the addition reaction of PEG
vinylsulfones and cysteine containing oligopeptides are affected by the branching
factor [82] and the molecular weight of the PEG macromer [74]. Additionally, the
final network properties depend on the precursor concentration and the stoichiometry
of reactive groups [74].
The network properties and swelling characteristics are further related to the mass
transport characteristics of hydrogels [9, 10, 83, 84]. To accomplish a time-delayed
release of small organic drugs or growth factors, for instance, it is necessary to limit
the free diffusion out of the hydrogel carrier [9, 10, 85]. On the other hand, enhancing
the supply of oxygen and nutrients as well as the removal of waste products is essential
for the survival and growth of the implanted cells [83, 84, 86].
Besides appropriate mechanical properties and mass transport characteristics,
degradation of the hydrogel is essential for many tissue engineering applications.
Admittedly, most hydrogels formed by cross-linking of macromers exhibit a strong
interdependency of cross-link density, mechanical properties, and degradation rate.
With regard to the desired characteristics of the hydrogel, however, the independent
control of degradation rate and mechanical properties will be crucial.
Ionically cross-linked hydrogels, such as alginate gels, normally undergo slow disso-
lution due to complexation of divalent cations or gradual exchange with monovalent
cations present in the environment [78]. Reducing the molecular weight of alginate
polymer chains [87] and introduction of hydrolytically labile acetal-like groups by
oxidation [88, 89] allows for control of the degradation rate and mechanical properties
in an almost independent manner. Hydrogels formed by photo-polymerization of PEG
diacrylate or PEG dimethacrylate are non-degradable within the typical timescale of
cell culture experiments. To render these hydrogels bioerodible, poly(α-hydroxy es-
ters), such as poly(lactic acid) (PLA) or poly(glycolic acid) (PGA), have been grafted
27
Chapter 2 Rational design of hydrogels for tissue engineering
to the PEG central block finally leading to triblock copolymers (PLA-b-PEG-b-PLA
or PGA-b-PEG-b-PGA) with acrylate or methacrylate end groups. The degradation
rate can be tailored by appropriate choice of the hydrolyzable poly(α-hydroxy esters)
and by varying its block length [72]. Cross-linking of PEG vinylsulfone macromers
with enzymatically cleavable peptides, such as matrix metalloproteinase (MMP) sen-
sitive peptides, allows for the creation of hydrogels that are susceptible to proteolytic
breakdown. The degradation kinetics were found to depend on the MMP activity of
the incorporated substrate and the action of cell-secreted MMPs [90].
2.5 Physical parameters regulate tissue developmentin vitro and in vivo
Current research efforts focus on physical cues regulating cell function and tissue
morphogenesis. Therefore, the physical characteristics of biomaterials used in tissue
engineering applications should no longer be neglected with respect to their biological
effects [22–24]. The subsequent chapters are to illustrate the impact of substrate
stiffness and degradability on tissue engineering. Thereby, we will focus on the use of
hydrogels and outline the effects of their inherent properties on tissue morphogenesis.
The effects of externally applied forces on cells and tissues are reviewed elsewhere in
detail [22, 24, 91, 92] and will, therefore, not be addressed here.
2.5.1 Impact of mechanical factors on cell function and tissuemorphogenesis
In order to assess the impact of hydrogel pore size on neurite extension, Dillon et al.
entrapped dorsal root ganglions (DRGs) into agarose gels of varying concentration.
Concomitantly with increasing agarose concentration, the average pore size decreased
exponentially as calculated from hydraulic permeability measurements. Similarly,
the length of extended neurites decreased with increasing agarose concentration [93].
In a follow-up study, Balgude et al. correlated the rate of neurite extension to
the mechanical stiffness of the hydrogel. They prepared agarose gels of varying
28
2.5 Physical parameters regulate tissue development
concentration and determined G∗ by oscillatory rheometry. The magnitude of G∗ was
used to calculate the force exerted by the hydrogel network on the advancing neurite
growth cones. Thereby, Balgude et al. found an inversely proportional relationship
between the force exerted by the hydrogel and the rate of neurite extension [94].
Similar results were obtained by Gunn et al. who encapsulated PC12 cells, a
commercially available rat pheochromocytoma cell line, into photo-cross-linkable
hydrogels prepared from PEG diacrylate. The Young’s modulus significantly increased
when the weight fraction of PEG diacrylate was increased. To mediate cell attachment,
hydrogels were further functionalized with various adhesion ligands. As a result of
this study, neurite extension was found to depend on the type and concentration
of adhesion ligand as well as the mechanical properties of the hydrogel. Compared
to more flexible hydrogels, gels with higher modulus significantly decreased neurite
extension [79].
To investigate the influence of cross-link density on cartilaginous tissue formation,
Bryant et al. embedded bovine chondrocytes into hydrogels prepared from PEG
dimethacrylate. Swelling studies revealed an increase in cross-link density with in-
creasing macromer concentration. After cultivation, immunohistochemistry suggested
an enhanced production of collagen type II in hydrogels of intermediate cross-link
density. Deposited collagens and glycosaminoglycans (GAGs) were primarily lo-
cated pericellularly, indicating that diffusion of macromolecules is restricted within
these gels. Only in the most loosely cross-linked hydrogels GAGs were distributed
homogenously [80].
Cartilaginous tissue formation was also studied by Wong et al. using alginate
hydrogels. In this study, the alginate type was shown to affect ECM accumulation,
whereby gels containing intermediate amounts of guluronic acid showed the highest
level of matrix synthesis. Among other possible reasons, such as impurities of the
different alginate types, ECM production is thought to be influenced by the mechanical
stiffness of the hydrogel, which results from the alginate type utilized [95].
Capillary morphogenesis has also been shown to depend on the substrate stiffness.
Sieminski et al. cultured human blood outgrowth endothelial cells (HBOECs) and
human umbilical vein endothelial cells (HUVECs) in collagen gels that were either
free floating or bound to the bottom of the well. The apparent stiffness of the matrix
is thought to depend on the collagen concentration as well as whether the gels are free
29
Chapter 2 Rational design of hydrogels for tissue engineering
floating or attached to the rigid culture plastic. Generally, capillary morphogenesis
seemed to be improved in more malleable environments. Furthermore, the apparent
matrix stiffness that supported capillary morphogenesis to the highest extent was
found to vary with different endothelial cells and their ability to contract the collagen
matrix [96].
These examples illustrate the impact of mechanical cues on cell behavior and tissue
morphogenesis. Cells embedded into hydrogels probably sense some sort of physical
confinement that regulates growth, differentiation, and ECM accumulation. This
confinement may be caused by the mechanical properties of the hydrogel itself as well
as the pericellular accumulation of ECM macromolecules. The supply of nutrients,
oxygen, and bioactive substances, as well as the removal of waste products, are also
affected by the network properties and swelling characteristics of hydrogels [9, 10, 83,
84, 86]. This, in turn, may also contribute to the observed cellular responses.
2.5.2 Influence of degradation profile on tissue formation
As outlined above, tissue morphogenesis is strongly influenced by the mechanical
properties of the supportive matrix. However, as most biomaterials used for tissue
engineering applications are biodegradable, the initial mechanical properties are not
retained over time. During the degradation of hydrogels, the average mesh size and
swelling level increase, and the diffusion of macromolecules, e.g. ECM components, is
facilitated. Concomitantly with the increase in mesh size, the mechanical properties
of the degrading hydrogel decrease significantly.
To examine the effects of temporally changing physicochemical properties on tissue
formation, Bryant et al. encapsulated bovine chondrocytes into photo-cross-linkable
hydrogels prepared from varying ratios of degradable PLA-b-PEG-b-PLA diacrylate
and nondegradable PEG dimethacrylate. After six weeks of cultivation, the total
collagen and desoxyribonucleic acid (DNA) contents were significantly increased in
gels with a high proportion of degradable macromers. The synthesis of collagen
type II also seemed to be favored, as indicated by immunohistochemistry. Altogether,
in highly degradable hydrogels, the composition of deposited ECM (collagens and
GAGs) more closely approached those of native cartilage, compared to less degradable
30
2.5 Physical parameters regulate tissue development
gels. Additionally, the secreted ECM was distributed more homogeneously throughout
the whole tissue, whereas, in gels with less degradable macromers, ECM was mainly
located in the pericellular region [97].
Alsberg et al. compared irradiated, more rapidly degrading alginate hydrogels
and non-irradiated, slowly degrading gels regarding their ability to support bone
development in vivo. Rat-derived osteoblasts were encapsulated into RGD-modified,
calcium cross-linked alginate gels and implanted into the backs of mice. Histological
examinations, bone densitometry, and microcomputed tomography (µCT) revealed
that rapidly degrading gels dramatically improved the extent and quality of bone
formation [87].
Similar results can be found by Kong et al., who used non-oxidized, high MW
alginates and binary blends of oxidized, low and high MW alginates. Rat-derived
BMSCs embedded in RGD-conjugated, calcium cross-linked alginate gels were im-
planted in the backs of mice. In order to promote differentiation of the BMSCs to
osteoblasts, the hydrogels were loaded with bone morphogenic protein-2 (BMP-2) and
transforming growth factor-β3 (TGF-β3). Compared to the more slowly degrading
non-oxidized, high MW gels, the more rapidly degrading binary gels facilitated the
formation of new bone tissue, indicated by histological sections [88].
However, tailoring the degradation rate not only provides control over tissue
morphogenesis. In a recently published work, Mahoney et al. reported the temporal
control of neural tissue formation by altering the degradation rate of methacrylate
end-capped triblock copolymers of PLA, PGA, and PEG. During the first week
of culture, photoencapsulated neural cells (precursor cells and neurons) assembled
together and formed small micro-tissues, which are considered to be building blocks
for the creation of functional neural circuits. After two weeks, the mesh size of the
hydrogel exceeded a critical value and processes emerged to penetrate throughout
the environment. Immunocytochemistry further revealed the presence of neurons and
glial cells that were responsive to neurotransmitters. As the time-scale over which
neural tissue develops could be tailored by incorporation of the cells into quickly
degrading (PGA-b-PEG-b-PGA) or more slowly degrading networks (PLA-b-PEG-b-
PLA), Mahoney et al. identified the degradation rate as a critical factor influencing
process outgrowth and neural cell differentiation [98].
31
Chapter 2 Rational design of hydrogels for tissue engineering
Degradability and degradation rate of the supportive matrix were identified as having
a strong influence on cell migration, proliferation, differentiation, and morphology of
the newly formed tissue. But the examples outlined above also illustrate that it may be
hard to distinguish between effects of substrate degradability and substrate mechanics.
The observed biological responses may be due to the given mechanical properties of the
matrix or to the ongoing loss of material during degradation. Together, the presented
studies imply that cell differentiation and tissue morphogenesis are supported by
rapidly degrading matrices. This, however, may be a false conclusion, as the optimal
degradation rate will depend on the intended application as well as the specifics
of particular cells. The study of Meinel et al. illustrates this issue: too rapid of a
degradation rate caused collagen scaffolds to collapse before substantial amounts of
ECM were deposited by the cells [99]. Consequently, it would be beneficial to couple
the rate of matrix degradation to the rate of ECM production in order to support
cell differentiation and tissue integrity.
2.5.3 Cell-responsive hydrogels
Ideally, matrix degradation would occur in temporal and spatial synchrony with the
formation of new tissue. In traditional biomaterials, however, degradation typically
takes place by non-enzymatic cleavage of chemically labile bonds (e.g. by hydrolysis of
ester bonds). Therefore, adapting the degradation rate to the rate of tissue formation
is a challenging task. In contrast, cell-responsive biomaterials mimic the proteolytic
recognition of natural ECMs and are degraded by cell-secreted and cell-activated
proteases, such as MMPs and serine proteases. This creates a dynamic balance
between matrix degradation and ECM deposition and allows for the remodeling of
the biomaterial by encapsulated or invading cells [8, 15–17].
To study the invasion characteristics of human fibroblasts in vitro, Lutolf et al.
attached integrin-binding domains (RGDSP) to vinylsulfone-functionalized PEGs
and cross-linked the macromers with MMP-sensitive peptide sequences (Figure 2.6).
The cell invasion rate was found to depend on the RGD ligand density, the MMP-
sensitivity, and the cross-link density of the networks. At a constant RGD ligand
32
2.5 Physical parameters regulate tissue development
density, lowering the PEG molecular weight from 20 to 15 kDa decreased the invasion
rate by almost a factor of four [90].
Adhesion motifs Protease-sensitive peptides Invading cell
Figure 2.6: Michael-type addition reaction between four-armed PEG-vinylsulfones andcysteine containing peptides. Introduction of adhesion sequences (step 1). Gelation isperformed in the presence of cells using protease-sensitive peptides as cross-linker (step 2).Networks are locally degraded by cellular proteases (step 3). Reprinted with permissionfrom Lutolf et al. [90]. c© 2003 National Academy of Sciences U.S.A.
To demonstrate the suitability of these biomaterials to induce tissue regeneration in
vivo, similar gels were used to deliver physically entrapped BMP-2 to the site of critical-
sized defects in rat crania. After five weeks, bone healing was assessed by analyzing
cranial samples using µCT. MMP-sensitive, BMP-2-bearing hydrogels promoted bone
formation to a significantly higher extent than MMP-insensitive or BMP-2-lacking
controls. Regarding the bone coverage, no significant differences could be observed
between hydrogels composed of four-armed PEGs of 15 and 20 kDa molecular weight.
However, cell infiltration was much more extensive in gels of lower cross-link density
(higher MW), resulting in increased bone volume and connectivity [100].
In another study, Park et al. evaluated the suitability of MMP-sensitive PEG-based
hydrogels as scaffolds for cartilage repair. Bovine chondrocytes were cultured for one
month in hydrogels prepared from PEG-vinylsulfones and MMP-sensitive peptides
(degradable gels) or PEG-vinylsulfones and PEG-dithiols (non-degradable gels). When
eight-armed PEG-vinylsulfone is used instead of four-armed, the cross-linked hydrogels
are stiffer and less swollen. The increased stiffness not only significantly reduced
cell number and size of the cultured clusters, but also influenced the expression of
ECM genes and the spatial distribution of deposited ECM. In degradable hydrogels,
the expression level of collagen type II, collagen type XI, and aggrecan increased
33
Chapter 2 Rational design of hydrogels for tissue engineering
with decreasing cross-link density, indicated by real-time polymerase chain reaction
(PCR), and ECM fibrils extended far into the hydrogel. In contrast, non-degradable
hydrogels generally led to lower expression levels of ECM genes and densely packed
ECM fibrils, but supported the expression of MMP-13 genes. Due to these results,
Park et al. concluded that cells obviously sense the matrix density or different levels
of physical confinement and subsequently adapt their gene expression pattern [82].
The degradation rate of these cell-responsive hydrogels depends on the action
of cellular proteases rather than environmental factors, such as the pH. Therefore,
the mechanical properties of the gels are thought to be retained until cell-triggered
remodeling is initiated. This enables the fabrication of hydrogels that provide an
appropriate mechanical environment for cell migration, proliferation, and differentia-
tion throughout their lifetime. The sustained mechanical integrity also prevents early
failure of the scaffold in load bearing applications. Taken together, these hydrogel
systems allow for an enormous versatility in design and might be useful in a variety
of tissue engineering applications.
2.6 Concluding remarks
Physical factors are potent regulators of cell function and tissue morphogenesis. In
most cases, however, the observed biological effects cannot be attributed to individual
physical parameters: the network properties and swelling characteristics of hydrogels,
for example, determine their mechanical properties as well as their mass transport
characteristics. Consequently, the diffusivity of growth factors, oxygen, nutrients, or
waste products will also contribute to the cellular response. In addition, the initial
mechanical properties of biomaterials are not the only factors decisive for the cellular
fate: over time, the substrate mechanics may significantly change due to degradation
of the matrix and/or deposition of ECM macromolecules. To survey this process,
non-invasive and non-destructive methods of measuring the mechanical properties are
required. However, emerging techniques, such as AFM and MRE, are still far from
being routine in tissue engineering. Therefore, the application of these methods has
to be further propagated in order to allow for the systematic characterization of living
tissues. Modeling of cell-biomaterial interactions in three-dimensional matrices may
34
2.6 Concluding remarks
also contribute to understand the complex interdependency of substrate mechanics
and adhesiveness [19–21, 101]. Together, this enables the integration of chemical,
physical, and topographical aspects and may have a powerful impact on the rational
design of appropriate biomaterials for tissue engineering applications.
35
Chapter 3
Poly(ethylene glycol) based hydrogelsfor intraocular applications
Ferdinand Brandl1, Matthias Henke1, Stefan Rothschenk1,2,
Ruth Gschwind3, Miriam Breunig1, Torsten Blunk1, Jorg Teßmar1,
Achim Gopferich1
1 Department of Pharmaceutical Technology, University of Regensburg, 93040 Regensburg2 LTS Lohmann Therapie-Systeme AG, 56626 Andernach3 Department of Organic Chemistry, University of Regensburg, 93040 Regensburg
Published in Advanced Engineering Materials 9 (12), 1141–1149 (2007).
37
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
Abstract
Hydrogels are attractive materials for biomedical applications due to their versa-
tility and excellent biocompatibility. In this study, we report the preparation of
poly(ethylene glycol) (PEG) based hydrogels for intraocular applications. We syn-
thesized branched PEG-succinimidyl propionates (10 kDa molecular weight) and
different types of PEG-amines (linear and branched, 2 and 10 kDa molecular weight).
Transparent hydrogels were formed in situ upon chemical reaction of these macromers.
The gels were characterized by oscillatory rheometry and NMR experiments. By
varying the concentration of macromers, the functionality of the PEG-amine, and
the conditions during cross-linking, gels with adequate gelation times of approx. 5 –
10 min and gel strengths of approx. 350 – 1500 Pa were obtained. The cross-linked
hydrogels showed no cytotoxic effects and may be used as vitreous substitutes or
intraocular drug release systems.
38
3.1 Introduction
3.1 Introduction
Hydrogels are highly hydrated networks of interacting polymer chains with viscoelastic
properties similar to those of natural tissues. Due to their enormous versatility and
excellent biocompatibility, they have been used for a variety of biomedical applications.
In tissue engineering approaches, hydrogels were studied as cell carriers for the
regeneration of a wide range of tissues including bone, cartilage, muscle, and neuronal
tissue. They were also applied as controlled release systems for bioactive molecules
(e.g. growth factors, nucleic acids, and various drugs) or as space filling scaffolds in
plastic surgery [7, 8, 10, 102, 103].
When designing new biomaterials for such sophisticated applications, an exact
knowledge of the biochemical and physicochemical requirements is essential. Hydrogels
designed for intraocular applications, for example, need to have distinct optical and
mechanical characteristics that are close to the properties of the vitreous body. The
natural vitreous is a transparent, virtually acellular, gel-like network of collagen fibrils
and glycosaminoglycans that fills the posterior segment of the eye. It acts as a shock
absorber, maintains the shape of the eye, and assists in holding the neuronal retina
in place [104, 105]. Dysfunctionalities of the vitreous body due to aging, traumatic
injuries, tumors, or systemic diseases (e.g. diabetes mellitus) often result in severe
visual impairment [105–107]. Proliferative diabetic retinopathy (PDR), for example,
is a frequently occurring complication of diabetes mellitus and characterized by an
abnormal growth of blood vessels into the vitreous body [108]. Besides age-related
macular degeneration (AMD), cataract, and glaucoma, PDR is one of the leading
causes of blindness in industrialized nations [18].
In such cases, a total replacement of the affected vitreous may be required in order
to prevent blindness or to restore vision [105–107]. However, today’s clinically used
substitutes (e.g. silicone oil, perfluorocarbon liquids, and gases) differ significantly
from the natural vitreous body regarding their physicochemical properties and me-
chanics. Some of these materials are also associated with severe side-effects when
kept intravitreally over longer periods of time [105, 106]. As alternatives, various gel-
forming polymers (e.g. collagen, polysaccharides, and synthetic polymers) have been
studied over the past decades, but none of them met clinical standards due to rapid
degradation, fast clearance from the eye, and numerous other complications [105–107].
39
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
Cross-linked hydrogels, which are expected to exhibit longer retention times, also
failed as vitreous substitutes as their mechanical properties are seriously impaired
upon the injection process [107, 109]. It has been hypothesized that these difficulties
could be avoided if low-viscous gels were used that solidify after injection into the
vitreous cavity. Apart from their potential as vitreous substitutes, such hydrogels
would be promising materials for the delivery of drugs to the posterior segment of
the eye. As the pharmacologic treatment of vitreoretinal diseases, such as PDR and
AMD, fairly advanced during the last years, therapeutic benefits can be expected
from hydrogels loaded with anti-inflammatory or anti-proliferative agents [110, 111].
Hydrogels based on poly(ethylene glycol) (PEG) can be considered potential vitreous
substitutes due to their excellent biocompatibility and transparency. Aqueous solutions
of PEG macromers can be cross-linked in situ, if chemical reactions are used that
are sufficiently gentle to be performed in the presence of cells or in vivo [73]. First
studies that can be found in the literature are highly promising. Glucose-permeable
hydrogels have been developed by cross-linking star-shaped PEG-amines with a di-
succinimidyl ester of PEG. The gels formed in water without any catalysts or initiators
and showed good biocompatibility when implanted subcutaneously in rats [112]. A
rapidly gelling tissue sealant based on thiol-functionalized PEG macromers and PEG-
succinimidyl glutarates has also been described [113]. Lutolf et al. prepared hydrogels
by stepwise copolymerization of vinylsulfone-functionalized PEG macromers and
cysteine containing peptides that were successfully applied in a variety of tissue
engineering applications [74, 82, 100]. For ophthalmologic applications, hydrogels
consisting of lysine-terminated dendrons and PEG-succinimidyl propionates have
been proposed for the closure of scleral incisions [114].
Despite their promise for other applications, the described hydrogels would be less
suited for intraocular applications due to their high polymer content of up to 40 % (w/v)
and their high mechanical stiffness of several thousand Pascal. Furthermore, most of
these gels solidify within a few seconds, which would impede their injection into the
eye. In this paper, we report the synthesis and preparation of hydrogels cross-linked
in situ by chemical reaction of branched PEG-succinimidyl propionates with PEG-
amines. As the two macromers are linked together by amide bonds, the obtained
hydrogels are expected to be stable over an extended period of time. Gelation kinetics
and mechanical strength were analyzed by oscillatory shear experiments and adjusted
40
3.2 Materials and Methods
to the requirements of potential vitreous substitutes. The effectiveness of cross-linking
was further evaluated by nuclear magnetic resonance (NMR) experiments. In vitro
cytotoxicity of the cross-linked gels was investigated using a standard cell proliferation
assay. In a final experiment, we studied the suitability of the prepared hydrogels as
potential drug carrier systems which would be interesting for the pharmacotherapy of
common eye diseases, such as PDR and AMD.
3.2 Materials and Methods
3.2.1 Materials
Acrylonitrile, N,N’ -dicyclohexylcarbodiimide (DCC), diisopropyl azodicarboxylate
(DIAD), Dulbecco’s modified Eagle’s medium (DMEM), fetal bovine serum (FBS), fluo-
resceine isothiocyanate (FITC)-dextrans, N-hydroxysuccinimide (NHS), poly(ethylene
glycol) (molecular weight 2 kDa, PEG2k-OH), and sodium dodecyl sulfate (SDS)
were purchased from Sigma-Aldrich (Taufkirchen, Germany). Phthalimide was ob-
tained from Acros Organics (Geel, Belgium). Four-armed poly(ethylene glycol)
(molecular weight 10 kDa, 4armPEG10k-OH) was purchased from Nektar Thera-
peutics (Huntsville, AL). Diethyl ether, ethanol, and methylene chloride were of
technical grade and used without purification. Deuterated chloroform (CDCl3) and
deuterium oxide (D2O) were purchased from Deutero GmbH (Kastellaun, Germany).
FluoSpheres R© and phosphate buffered saline (PBS) were purchased from Invitrogen
GmbH (Karlsruhe, Germany). All other chemicals were of analytical grade and
purchased from Merck KGaA (Darmstadt, Germany). Water was obtained using a
Milli-Q water purification system from Millipore (Schwalbach, Germany).
3.2.2 Synthesis of PEG-amines
PEG-amines were synthesized according to a procedure described by Mongondry et
al. [115]. For a typical synthesis, PEG2k-OH (5 mmol), phthalimide (15 mmol), and
triphenylphosphine (PPh3) (15 mmol) were dissolved in 50 mL of tetrahydrofuran
41
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
(THF). Then, a solution of DIAD (15 mmol) in THF (15 mL) was added dropwise
(Figure 3.1A). For the synthesis of branched PEG-amines, 4armPEG10k-OH (1 mmol)
was used instead of PEG2k-OH and the excess of reagents was decreased to 1.2 times.
After 48 h of stirring at room temperature, the solvent was evaporated under reduced
pressure. For purification, the raw product was dissolved in water again, filtered, and
washed with diethyl ether. After the water had been removed under reduced pressure,
the oily product was dissolved in methylene chloride and subsequently precipitated
by dropping into ice cold diethyl ether. The precipitate was collected by filtration
and dried under vacuum overnight.
The obtained phthalimido-PEGs (4 mmol) were dissolved in ethanol (50 mL) and
treated with hydrazine monohydrate (40 mmol) under reflux for 5 h. After cooling to
room temperature, hydrochloric acid (3 M) was added up to pH 2 – 3 and the formed
precipitate was removed by filtration. The solvent was evaporated, the obtained raw
product was dissolved in water, and the pH of the aqueous solution was adjusted with
sodium hydroxide (3 M) to 9 – 10. The product was then extracted with methylene
chloride. After drying over anhydrous sodium sulfate, the solution was concentrated
under reduced pressure and dropped into ice cold diethyl ether. The precipitated
polymer was collected and dried under vacuum.1H-NMR (PEG2k-NH2, CDCl3, 300 MHz): δ 2.95 ppm (t, 4H, –OCH2CH2NH2),
3.65 ppm (s, –OCH2CH2O–). 1H-NMR (4armPEG10k-NH2, CDCl3, 600 MHz):
δ 2.87 ppm (t, 8H, –OCH2CH2NH2), 3.41 ppm (s, 8H, R3CCH2O–), 3.64 ppm
(s, –OCH2CH2O–). The degree of end-group conversion ranged from 95 to 100 %, as
determined by NMR.
3.2.3 Synthesis of branched PEG-succinimidyl propionates
PEG-propionic acid was synthesized as described by Sedaghat-Herati et al. [116].
For the synthesis of branched succinimidyl esters (Figure 3.1B), the experimental
procedure was modified as follows: 4armPEG10k-OH (1 mmol) was dissolved in water
(15 mL) containing potassium hydroxide (0.20 g). After cooling to 0 ◦C, acrylonitrile
(20 mmol) was added stepwise. The reaction mixture was stirred for 2.5 h at 0 ◦C
and then kept in the refrigerator overnight. After neutralizing with hydrochloric
42
3.2 Materials and Methods
O O
O
O
OOH
OOH nn
OOH
OOH
n
n
RO
OO
CNn
RO
OO n OH
O
RO
OO n O
O
N
O
O
H C=CH–CN, KOH2
1. H O , KOH
2. KOH conc.2 2
NHS, DCC
A) B)
OO
OHOH n
OO
NN n
O
O
O
O
OO
NH2NH
2 n
PPh , DIAD,
phthalimide3
H N–NH2 2
Figure 3.1: Synthesis of PEG2k-NH2 (A) and 4armPEG10k-SPA (B).
acid (3 M), the aqueous solution was extracted with methylene chloride. The combined
organic phases were washed with a saturated solution of sodium chloride and dried
over anhydrous sodium sulfate. The organic solution was concentrated under reduced
pressure and dropped into ice cold diethyl ether. The precipitated product was
collected by filtration and dried under vacuum.
To obtain the PEG-propionic acid, PEG-propionitrile was dissolved in water (50 mL)
containing potassium hydroxide (0.18 g). This solution was cooled to 0 ◦C and 2.70 mL
of a hydrogen peroxide solution (30 %) were added stepwise. The reaction mixture was
stirred at room temperature overnight. Then, 1.5 g of sodium thiosulfate and 250 mL
of a potassium hydroxide solution (6 %) were added. After stirring for 48 h at room
temperature, the reaction mixture was placed in an ice bath and neutralized with
hydrochloric acid (3 M). The product was extracted and precipitated as described for
PEG-propionitrile.
For the synthesis of the corresponding succinimidyl ester, PEG-propionic acid
(0.2 mmol) and NHS (1.6 mmol) were dissolved in 15 mL of dry methylene chloride.
DCC (1.6 mL) was separately dissolved in 5 mL of the same solvent. The carbodiimide
solution was then added to the solution of PEG-propionic acid and kept under stirring
at room temperature overnight. The mixture was filtrated to remove precipitated
43
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
dicyclohexylurea, concentrated under reduced pressure and dropped into ice cold
diethyl ether. The precipitated polymer was collected and dried under vacuum.1H-NMR (4armPEG10k-SPA, CDCl3, 600 MHz): δ 2.84 ppm (s, 14H, –CH2CH2-
COOSu), 2.90 ppm (t, 7H, –CH2CH2COOSu), 3.41 ppm (s, 8H, R3CCH2O–), 3.64 ppm
(s, –OCH2CH2O–), 3.85 ppm (t, 7H, –CH2CH2COOSu). The degree of substitution
was about 90 %, with the remaining 10 % being PEG-propionic acid, as determined
by NMR spectroscopy.
3.2.4 Preparation and rheological characterization of hydrogels
Gelation kinetics and gel strength were studied by performing oscillatory shear experi-
ments on a TA Instruments AR 2000 rheometer (TA Instruments, Eschborn, Germany)
with parallel plate geometry under humidified atmosphere. For the preparation of
hydrogels, specified amounts of PEG-amines were dissolved in buffer solutions of
defined pH (Table 3.1); 4armPEG10k-SPA was dissolved in PBS or water, respectively.
The solutions of 4armPEG10k-SPA were filtrated (0.20 µm syringe filter, Corning
Costar, Bodenheim, Germany) to remove traces of water-insoluble impurities. The
two precursor solutions were mixed and immediately cast onto the lower plate of the
rheometer.
The evolution of storage (G′) and loss moduli (G′′) was recorded as a function of
time at designated temperatures (Table 3.1), 1 Hz oscillatory frequency, and 10 µNm
torque (20 mm steel plate, measuring gap size 1000 µm). The crossover of G′ and
G′′ was regarded as the gel point. The gel strength was determined by recording the
complex shear modulus (G∗) at a constant frequency of 1 Hz as a function of the
applied stress. If measurements were affected by the existent system inertia, G′ and
G′′ were recorded at 0.5 Hz oscillatory frequency and 10 µNm torque (40 mm steel
plate, measuring gap size 500 µm).
3.2.5 Characterization of hydrogels by NMR
For NMR measurements, 4armPEG10k-SPA and PEG2k-NH2 were dissolved in D2O
in concentrations of 10 % (w/v) and 4 % (w/v), respectively. Samples were prepared
44
3.2 Materials and Methods
Table 3.1: Composition of hydrogels and conditions of gel preparation
4armPEG10k-SPA PEG-amineGel Concentration Concentration Type pH Temperature Ratio ra
1 5 % (w/v) 1 % (w/v) linear (2 kDa) 7.4b 25 ◦C 2.00
2 5 % (w/v) 1.5 % (w/v) linear (2 kDa) 7.4b 25 ◦C 1.33
3 5 % (w/v) 2 % (w/v) linear (2 kDa) 7.4b 25 ◦C 1.00
4 5 % (w/v) 2.5 % (w/v) linear (2 kDa) 7.4b 25 ◦C 0.80
5 10 % (w/v) 3 % (w/v) linear (2 kDa) 7.4b 25 ◦C 1.33
6 10 % (w/v) 4 % (w/v) linear (2 kDa) 7.4b 25 ◦C 1.00
7 5 % (w/v) 1.5 % (w/v) linear (2 kDa) 6.0c 25 ◦C 1.33
8 5 % (w/v) 1.5 % (w/v) linear (2 kDa) 6.5c 25 ◦C 1.33
9 5 % (w/v) 1.5 % (w/v) linear (2 kDa) 7.0c 25 ◦C 1.33
10 5 % (w/v) 1.5 % (w/v) linear (2 kDa) 7.5c 25 ◦C 1.33
11 5 % (w/v) 2 % (w/v) linear (2 kDa) 7.0d 15 ◦C 1.00
12 5 % (w/v) 2 % (w/v) linear (2 kDa) 7.0d 25 ◦C 1.00
13 5 % (w/v) 2 % (w/v) linear (2 kDa) 7.0d 35 ◦C 1.00
14 2 % (w/v) 2 % (w/v) 4arm (10 kDa) 7.0d 25 ◦C 1.00
15 3 % (w/v) 3 % (w/v) 4arm (10 kDa) 7.0d 25 ◦C 1.00
aThe stoichiometric ratio r is defined as the molar ratio of succinimidyl ester to amino groups.Calculations are performed assuming quantitative conversion of end-groups.
bPhosphate buffered saline (both macromers dissolved in PBS)c250 mM phosphate buffer (PEG-amine in 500 mM phosphate buffer, 4armPEG10k-SPA in water)d25 mM phosphate buffer (PEG-amine in 50 mM phosphate buffer, 4armPEG10k-SPA in water)
from equal volumes of the filtrated precursor solutions. 1H-NMR and 13C-NMR
spectra were recorded at 600 and 150 MHz, respectively, on a Bruker Avance 600
(Bruker BioSpin GmbH, Rheinstetten, Germany). To follow the cross-linking of
hydrogels, a time series of 1H-NMR spectra was recorded over a total time of 180 min.
3.2.6 Cytotoxicity of cross-linked hydrogels
NIH 3T3 fibroblasts (LGC Promochem GmbH, Wesel, Germany) were grown in T-75
cell culture flasks containing 10 mL DMEM supplemented with 10 % FBS at standard
cell culture conditions (37 ◦C, 95 % relative humidity, and 5 % CO2).
45
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
The cytotoxicity of the cross-linked hydrogels was evaluated by a medium extraction
test. 300 µL of hydrogels consisting of 2 % (w/v) 4armPEG10k-SPA and 2 % (w/v)
4armPEG10k-NH2 were cast into 24-well plates (surface area approx. 2 cm2). After
60 min of cross-linking, the gels were covered with 1 or 2 mL DMEM and incubated
under standard cell culture conditions (see above) for 24 h (surface area to fluid
volume 1 and 2 cm2/mL). Prior to cytotoxicity testing, the cells were replated at 90 %
confluency and seeded into 96-well plates at a density of 12,000 cells per well. Twenty
hours after seeding, the culture medium was replaced by the extraction medium
(100 µL/well), followed by incubation for 24 h. Cells treated with 0.01 % SDS served
as positive control.
Cell viability was evaluated using the CellTiter 96 R© AQueous One Solution Cell
Proliferation Assay (Promega GmbH, Mannheim, Germany). 20 µL of Cell Titer
96 R© AQueous One Solution containing the tetrazolium salt 3-(4,5-dimethylthiazol-2-
yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) were added
to each well. After 3 h of incubation under standard cell culture coconditions (see
above), the amount of soluble formazan produced by cellular reduction of MTS was
quantified by recording the absorption at 490 nm using a TitertekPlus Microplate
Reader (Friedrich S. Bartolomey Labortechnik, Rheinbach, Germany). Cell viability
was normalized to that of fibroblasts cultured in medium without extracts. All
experiments were carried out in triplicate and the results are expressed as means ±standard deviation.
3.2.7 Release of FITC-dextrans and fluorescent nanospheres
For release experiments, fluorescein isothiocyanate labeled dextrans (FITC-dextrans)
and orange fluorescent nanospheres (FluoSpheres R©) were used as model substances.
FITC-dextrans of 4 and 150 kDa molecular weight were dissolved in 50 mM phosphate
buffer pH 7.0 at 1 % (w/v). FluoSpheres R© (bead diameter 100 nm) were supplied as
aqueous suspensions containing 2 % solids. To prepare hydrogels, aliquots of these
three solutions were mixed with a 4 % (w/v) solution of 4armPEG10k-NH2 in 50 mM
phosphate buffer pH 7.0. Subsequently, an equal volume of a 4 % (w/v) solution of
4armPEG10k-SPA in water was added. The resulting mixture was vortexed; samples
46
3.3 Results and discussion
of 500 µL were then cast into glass vials (inner diameter 24 mm) and allowed to gel
for 60 min. In order to determine the in vitro release kinetics, samples (1.1 mm height
× 24 mm diameter) were covered with 5 mL of 50 mM phosphate buffered saline pH
7.4 (containing 0.025 % sodium azide) and maintained at 37 ◦C on an orbital shaker
(∼ 50 rpm). At specified time intervals, 400 µL of the supernatant were collected and
replaced with fresh buffer. The amounts of released FITC-dextrans (λex = 490 nm,
λem = 520 nm) and FluoSpheres R© (λex = 540 nm, λem = 560 nm) were determined
on a PerkinElmer LS 55 Fluorescence Spectrometer (PerkinElmer, Rodgau-Jugesheim,
Germany). All experiments were carried out in triplicate and the results are expressed
as means ± standard deviation.
3.3 Results and discussion
3.3.1 Preparation and rheological characterization of hydrogels
Hydrogels were prepared by step-growth polymerization of branched PEG-succinimidyl
propionates with PEG-amines. Figure 3.2 shows a typical rheogram for a gel formed
by reaction of 5 % (w/v) 4armPEG10k-SPA and 2 % (w/v) PEG2k-NH2 at pH 7.0
and 25 ◦C (Table 3.1, gel 12). Directly after mixing the two precursor solutions,
the sample behaved like a free-flowing liquid (G′′ > G′). This quality is of special
advantage, as it would enable the injection into the eye using small-gauge needles.
The cross-over of G′ and G′′ after approx. 10 min was regarded as the gel point. In the
course of time, cross-linking further proceeded as indicated by the steadily increasing
value of G′. After approx. 30 min, highly elastic gels had formed (G′ � G′′); the
value of G′ exceeded that of G′′ by several orders of magnitude. Mechanically, these
hydrogels would be suitable for vitreous substitution, as they are expected to assist
in holding the neuronal retina in place after the natural vitreous has failed. The
mechanical energy generated by external forces, such as rubbing or hitting, or by heart
beats and eye movements, would be stored and then dissipated slowly [105, 107, 109].
More importantly, the mechanical properties will be unaffected by the injection
process as cross-linking is performed in the vitreous cavity. Therefore, failure of
47
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
the substitute due to liquefaction, as observable for hydrogels cross-linked prior to
injection [107, 109], can be avoided.
0.0001
0.001
0.01
0.1
1
10
100
1000
0 10 20 30 40 50 60
Time (min)
G'
(Pa
)
0.0001
0.001
0.01
0.1
1
10
100
1000
G"
(Pa)
G'
G"
Figure 3.2: Rheogram of a gel formed by polymerization of 5 % (w/v) 4armPEG10k-SPAwith 2 % (w/v) PEG2k-NH2 at pH 7.0 and 25 ◦C (Table 3.1, gel 12). The measurementwas performed at 0.5 Hz oscillatory frequency using a 40 mm steel plate with 500 µmgap size.
As shown in Figure 3.3A, gelation was accelerated with decreasing stoichiometric
ratio r of reactive groups. At r = 2.00, solidification occurred after more than 90 min.
An exact value could not be determined, as the rheological measurements were run
over 90 min only. The highest cross-linking rate was observed at r = 0.80. Under
these conditions, the gel point was reached within 17 min. The stoichiometric ratio r
also affected the strength of the prepared hydrogels. With decreasing r, the stiffness
of the gels first increased, reached an optimum at r = 1.00, and then decreased again.
This is attributed to structural defects in the network architecture that occur upon
mixing of nonstoichiometric ratios of reactants [117]. At high stoichiometric ratios,
cross-linking will be inefficient due to the relative excess of succinimidyl ester groups.
In contrast, an excess of amino groups at low stoichiometric ratios will promote the
formation of cyclizations and elastically inactive dangling ends. Both phenomena
increase the average molecular weight between cross-links and lead to mechanically
weaker gels.
48
3.3 Results and discussion
10 % (w/v) 4armPEG10k-SPA(pH 7.4, 25 °C)
5 % (w/v) 4armPEG10k-SPA( = 1.00, pH 7.0)r
5 % (w/v) 4armPEG10k-SPA(pH 7.4, 25 °C)
5 % (w/v) 4armPEG10k-SPA( = 1.33, 25 °C)r
A)
D)C)
B)
Stoichiometric ratio r
2.00 1.33 1.00 0.80
Ge
lati
on
tim
e(m
in)
0
20
40
60
80
100
|G*|
(Pa
)
1
10
100
1000
10000
> 90
70
34
2017
295
460
165
Stoichiometric ratio r
1.33 1.00G
ela
tio
nti
me
(min
)0
20
40
60
80
100
|G*|
(Pa
)
1
10
100
1000
10000
13
8
16451390
pH
6.5 7.0 7.5
Ge
lati
on
tim
e(m
in)
0
20
40
60
80
100
|G*|
(Pa
)
1
10
100
1000
10000
23
9
1
200
355
1040
Temperature (°C)
15 25 35
Ge
lati
on
tim
e(m
in)
0
20
40
60
80
100
|G*|
(Pa
)
1
10
100
1000
10000
32
11
4
295
8401165
Figure 3.3: Dependence of gelation time (white bars, left y-axis) and gel strength (blackbars, right y-axis) on the stoichiometric ratio r of reactive groups, pH, and temperature.(A) Gels were formed from 5 % (w/v) 4armPEG10k-SPA and varying concentrations ofPEG10k-NH2 in PBS (pH 7.4) at 25 ◦C. (B) Gels were formed from 10 % 4armPEG10k-SPA and two different concentrations of PEG2k-NH2 in PBS at 25 ◦C. (C) Gels wereformed from 5 % (w/v) 4armPEG10k-SPA and 1.5 % (w/v) PEG2k-NH2 in 250 mM phos-phate buffer solutions at 25 ◦C. (D) Gels were formed from 5 % (w/v) 4armPEG10k-SPAand 2 % (w/v) PEG2k-NH2 at pH 7.0 (25 mM phosphate buffer). These measurementswere performed at 0.5 Hz oscillatory frequency using a 40 mm steel plate with 500 µmgap size.
49
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
Gelation time and gel strength also depended on the used concentrations of
macromers. Gels formed from 10 % (w/v) 4armPEG10k-SPA (Figure 3.3B) so-
lidified more than twice as fast as the corresponding gels containing 5 % (w/v)
4armPEG10k-SPA (Figure 3.3A). The mechanical strength was also dramatically
increased. These findings can again be explained by the architecture of the formed
polymer networks [117]. At low concentrations of 4armPEG10k-SPA, the overall con-
centration of succinimidyl ester groups is correspondingly low. The local concentration
of succinimidyl ester functions upon one PEG macromer, however, is independent
of the overall concentration. Thus, low concentrations of 4armPEG10k-SPA (Fig-
ure 3.3A) favor intramolecular cyclizations that decrease the cross-link density and
mechanical rigidity of the formed networks. In contrast, higher concentrations of
4armPEG10k-SPA (Figure 3.3B) favor intermolecular reactions as the probability
increases that cross-links are formed between two different 4armPEG macromers.
To investigate the influence of pH on the gelation rate, gels were made from solutions
of 4armPEG10k-SPA in water and solutions of PEG2k-NH2 in 500 mM phosphate
buffer solutions of defined pH (Figure 3.3C). Even though the ionic strength of these
buffers is far from being physiological, this high salt concentration was necessary to
ensure a constant pH during cross-linking. At pH 6.0, no gelation occurred within
180 min (data not shown). By increasing the pH from 6.5 to 7.5, the gelation time
was reduced from 23 min to approx. 1 min. Concomitantly, the stiffness increased by
a factor of 5. This pH dependency is attributed to the basic character of PEG2k-NH2
and the assumption that only free amine bases are able to react with succinimidyl
ester groups. Compared to pH 6.5, the amount of free amine bases is increased 10-fold
at pH 7.5. Obviously, this is sufficient to enhance the polymerization rate and the
resulting gel strength. Adjusting the pH is, therefore, seen as a valuable tool for
controlling the gel properties in future applications.
The cross-linking temperature was expected to be another important parameter
influencing gel properties. At 35 ◦C, the gel point was reached after approx. 4 min.
With decreasing temperature, the polymerization rate was reduced: at 25 and 15 ◦C
it took about 11 and 32 min, respectively, until gelation occurred (Figure 3.3D). At
15 ◦C, cross-linking was still incomplete even after 90 min (data not shown).
When comparing the gels formed in PBS (Figure 3.3A) with those prepared in
stronger buffer solutions (Figure 3.3C and D), significant differences became obvious.
50
3.3 Results and discussion
The former were mechanically weaker and showed lower polymerization rates than
the latter. This is associated with the fact that 4armPEG10k-SPA was dissolved in
PBS instead of water. As the hydrolysis rate of succinimidyl ester groups is higher in
PBS than in pure water (data not shown), some loss of 4armPEG10k-SPA during
sample preparation might explain the observed differences. Moreover, the pH was
found to slightly decrease during cross-linking, probably due to the acidic reaction of
released NHS (data not shown). Due to its low buffer capacity, PBS is not able to
compensate this drop in pH, which finally slows down the cross-linking rate.
To cope with the problem of inefficient cross-linking due to formation of intramolec-
ular cyclizations and elastically inactive dangling ends, hydrogels were formed upon
reaction of 4armPEG10k-SPA with branched PEG-amines. The overall concentra-
tion of macromers was 4 % (w/v) and 6 % (w/v), respectively. Due to the higher
functionality of 4armPEG10k-NH2 (4 vs. 2), the cross-linking rate of these hydro-
gels was increased compared to gels made from PEG2k-NH2. The gelation time
was approx. 5 min for gels containing 2 % (w/v) 4armPEG10k-NH2 and 2 min for
those containing 3 % (w/v). The stiffness reached values of about 1485 and 3220 Pa,
respectively. This high gel strength most likely originates from the reduced mesh size
of these networks (theoretically 5 kDa), which is lower than those of networks formed
from 4armPEG10k-SPA and PEG2k-NH2 (theoretically 7 kDa). Further, elastically
inactive cyclizations will not be generated if two branched macromers are used.
With respect to intraocular applications, hydrogels with an intermediate gelation
time are preferred, since these gels would be suitable for injection but solidify in an
appropriate period of time (approx. 5 – 10 min). This would be an essential feature of
potential vitreous substitutes, as it avoids their own drainage through retinal breaks
and ensures a proper position of the retina [105, 107]. Regarding the stiffness of
potential vitreous replacements, values similar to those of the natural vitreous body
would be desired. Defining theses values, however, might be a challenging task, as
the mechanical properties of the vitreous change with age, species, and preparation
method. The reported values of the storage modulus G′ ranged from a few Pascal for
porcine vitreous up to several hundred Pascal for goat vitreous [118, 119]. Indeed,
the stiffness of the developed hydrogels ranges at the upper limit of the reported
values but still can be adjusted by varying the gel composition and conditions during
cross-linking.
51
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
Due to their high water content of approx. 96 %, ideal gelation time, and appropriate
mechanical stability, hydrogels prepared from 2 % (w/v) 4armPEG10k-SPA and 2 %
(w/v) 4armPEG10k-NH2 were favored and used for cytotoxicity testing and release
experiments (see below).
3.3.2 Characterization of hydrogels by NMR
To investigate polymerization kinetics and cross-linking efficiency, a time series of1H-NMR spectra was recorded. The first spectrum was obtained approx. 5 min
after mixing the two precursor solutions and showed the characteristic signals of
4armPEG10k-SPA and PEG2k-NH2. The protons of the succinimidyl group (Fig-
ure 3.4, signal c) were found at 2.87 ppm. The methylene protons neighboring the
succinimidyl ester group (Figure 3.4, signal d) and amino group (Figure 3.4, signal e)
were identified at 2.95 and 3.13 ppm, respectively. The signal at 3.42 ppm (Figure 3.4,
signal g) corresponded to the pentaerythritol core of 4armPEG10k-SPA and was used
for the calibration of integrals. The methylene protons of the PEG backbone were
found at about 3.62 ppm (not shown in Figure 3.4). Moreover, 3 further signals
(Figure 3.4, signals a, b and f) appeared that were neither detected in the spectrum
of pure 4armPEG10k-SPA nor in that of pure PEG2k-NH2. The signal b at about
2.65 ppm stemmed from the methylene protons of released NHS; signals a and f at
2.46 and 3.32 ppm were assigned to the methylene protons neighboring the newly
formed amide bond. This was confirmed by HMBC spectra showing an interaction
between these protons and the carbonyl carbon atom at 174 ppm (data not shown).
Integration of the aforementioned signals revealed that a large fraction of succin-
imidyl ester groups had already reacted during preparation and acquisition of the
first spectrum. This, however, does not conflict with the rheological measurements,
as it is possible that in early stages of polymerization, weakly cross-linked polymer
strands or smaller aggregates are formed rather than percolating networks. Thus,
the cross-linking density might increase for a long time without enhancing the gel
strength significantly.
Within the observation period, the signals c, d and e decreased by approx. 30 %.
Concomitantly, the signals a, b and f increased by approx. 30 %, which is consistent
52
3.3 Results and discussion
CH
2
CH
2
NO O
OH
NH
O
C
H2
NH
CH
2
O
NH2
C
H2
O SuO CH
2
O
CH
2
CH
2
NO O
OR
CH
2
O
60 min
180 min
120 min
Start
2.402.502.602.702.802.903.003.103.203.30
ab
cde
fg
*
d (ppm)
3.40
Figure 3.4: 1H-NMR spectra of a hydrogel formed by reaction of 5 % (w/v) 4armPEG10k-SPA and 2 % (w/v) PEG2k-NH2 (D2O, 600 MHz, 300 K). The first spectrum (start)was obtained approx. 5 min after mixing the two precursor solutions. Then, spectra wererecorded for 180 min.
with the supposed mechanism of cross-linking. At later time-points, another signal
was recognized at 2.62 ppm that had been overlapped by signal b before (see asterisk in
Figure 3.4). This signal most likely corresponds to the PEG-propionic acid protons and
suggests occurring hydrolysis of the succinimidyl ester group. Furthermore, the integral
of signal e stayed somewhat higher than expected. This indicates a nonstoichiometric
consumption of amino groups probably due to hydrolysis of succinimidyl ester groups.
As a consequence, the cross-linked hydrogels contain small amounts of carboxylic acid
and ammonium groups; cross-linking efficiency and mechanical stability are, therefore,
somewhat lower than expected from theory. Altogether, NMR spectroscopy was found
to be a valuable method to monitor the process of gel formation on a molecular level.
3.3.3 Cytotoxicity of cross-linked hydrogels
Although the described hydrogels are being developed for intraocular applications,
this study was carried out with fibroblasts. Fibroblasts are widely accepted for
cytotoxicity testing [120] and used as a standard cell type in order to determine the
53
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
general biocompatibility of materials. To assess cytotoxicity, hydrogels consisting of
2 % (w/v) 4armPEG10k-SPA and 2 % (w/v) 4armPEG10k-NH2 were evaluated by a
medium extraction test. The cytotoxic effects of extracted products are presented
in Figure 3.5. The tested hydrogels showed no adverse effects on the viability of
fibroblasts, where the minimum viability was 98±4 %. Therefore, it can be concluded
that no toxic products (e.g. byproducts of the synthesis) are leached out of the
incubated gels. Detrimental effects due to shifts in pH or osmotic pressure can
be excluded. In future experiments, the biocompatibility with ocular tissues will
be investigated using a perfusion culture model of full-thickness porcine retina, an
in vitro model suitable for the evaluation of biomaterials in more organotypical
environments [121, 122].
Medium 0.01% SDS 1 cm²/ml 2 cm²/ml
Ce
llvia
bilit
y(%
)
0
20
40
60
80
100
120
Figure 3.5: Cell viability after exposure toextracts of hydrogels prepared from 2 %(w/v) 4armPEG10k-SPA and 2 % (w/v)4armPEG10k-NH2 (surface area to fluidvolume 1 and 2 cm2/mL). Data representmean ± standard deviation (n = 3 foreach group).
3.3.4 Release of FITC-dextrans and fluorescent nanospheres
The in vitro release properties of hydrogels were studied using FITC labeled dextrans
(molecular weight 4 and 150 kDa) and fluorescent nanospheres (FluoSpheres R©, bead
diameter 100 nm) as macromolecular model substances. Hydrogels were formed from
2 % (w/v) 4armPEG10k-SPA and 2 % (w/v) 4armPEG10k-NH2 (water content:
96 %). FITC-dextrans with a molecular weight of 4 kDa were released very rapidly,
indicating that the size of these molecules was smaller than the mesh size of the
prepared hydrogels (theoretically 5 kDa). After an initial burst, the release continued
54
3.3 Results and discussion
and was completed after approx. 10 h (Figure 3.6). The release exceeded 100 %
probably due to small measuring errors that propagate in the calculation of the
cumulative release data. However, this is not significant as it was the aim of this
study to characterize the overall release kinetics of model substances rather than
the absolute amounts released. The release rate of larger FITC-dextrans (molecular
weight 150 kDa) was considerably lower. Over a period of 10 h, the gels had released
approx. 70 % of the incorporated FITC-dextrans (Figure 3.6). Unlike smaller dextrans,
the release was completed only after 48 h. In contrast to the tested FITC-dextrans,
no significant release of FluoSpheres R© was observed within 96 h (data not shown).
The used FluoSpheres R© consist of polystyrene and have a bead diameter of 100 nm.
Most likely, they are trapped within the hydrogels and will not be released until the
mesh size increases due to degradation of the gels. In future studies, this could be
investigated using hydrogels prepared from biodegradable macromers.
Time (h)
0 2 4 6 8 10
Re
lea
se
dF
ITC
-de
xtr
an
(%)
0
20
40
60
80
100
120
140
FITC-dextran 4 kDa
FITC-dextran 150 kDa
Figure 3.6: Cumulative amount of FITC-dextrans (molecular weight 4 kDa and150 kDa) released from hydrogels consist-ing of 2 % (w/v) 4armPEG10k-SPA and2 % (w/v) 4armPEG10k-NH2. Data arerepresented as means ± standard devia-tions (n = 3 for each group).
These findings are of special interest regarding potential ophthalmologic applications
of the developed hydrogels. Similar to the natural vitreous body, the gels would allow
the diffusion of nutrients, metabolites, and soluble proteins to or from the adjacent
tissues. On the other hand, the hydrogels could be used as injectable carriers for
nanoparticulate drug delivery systems in order to treat common eye diseases such as
AMD or PDR, for example [110, 123].
55
Chapter 3 Poly(ethylene glycol) based hydrogels for intraocular applications
3.4 Conclusion
We successfully prepared optically transparent hydrogels by step-growth polymer-
ization of branched PEG-succinimidyl propionates with two different types of PEG-
amines. Gelation time and gel strength were found to depend on the macromer
concentration, the PEG-amine functionality, and the conditions during cross-linking.
By adjusting these parameters, we obtained hydrogels with ideal gelation times of
approx. 5 – 10 min and mechanical properties similar to those of the natural vitreous
body. The tested hydrogels showed no cytotoxic effects in vitro and may be used
as injectable vitreous substitutes. The gels allow for further functionalization by
incorporation of nanoparticulate drug delivery systems or by attachment of peptides,
proteins, and other bioactive substances, yielding biomimetic materials [124]. In
future applications, it would also be possible to load these hydrogels with cells natu-
rally occurring in the cortex of the vitreous body. These so called hyalocytes were
shown to accumulate collagen and hyaluronic acid in vitro and may be used in tissue
engineering approaches for the regeneration of the natural vitreous body [125].
56
Chapter 4
Hydrogel-based drug delivery systems:Comparison of drug diffusivity andrelease kinetics
Ferdinand Brandl1, Fritz Kastner2, Ruth Gschwind3, Torsten Blunk1,
Jorg Teßmar1, Achim Gopferich1
1 Department of Pharmaceutical Technology, University of Regensburg, 93040 Regensburg2 Center for Chemical Analysis, University of Regensburg, 93040 Regensburg, Germany3 Department of Organic Chemistry, University of Regensburg, 93049 Regensburg, Germany
Published in the Journal of Controlled Release, in press.
57
Chapter 4 Hydrogel-based drug delivery systems
Abstract
Hydrogels are extensively studied as matrices for the controlled release of macromole-
cules. To evaluate the mobility of embedded molecules, these drug delivery systems
are usually characterized by release studies. However, these experiments are time-
consuming and their reliability is often poor. In this study, gels were prepared by
step-growth polymerization of poly(ethylene glycol) (PEG) and loaded with fluores-
ceine isothiocyanate (FITC) labeled dextrans. Mechanical testing and swelling studies
allowed prediction of the expected FITC-dextran diffusivity. The translational diffu-
sion coefficients (D) of the incorporated FITC-dextrans were measured by fluorescence
recovery after photobleaching (FRAP) and pulsed field gradient NMR spectroscopy.
Because the determined values of D agreed well with those obtained from release
studies, mechanical testing, FRAP, and pulsed field gradient NMR spectroscopy are
proposed as alternatives to release experiments. The applied methods complemented
each other and represented the relative differences between the tested samples cor-
rectly. Measuring D can therefore be used to rapidly evaluate the potential of newly
developed drug delivery systems.
58
4.1 Introduction
4.1 Introduction
Hydrogels have been studied as injectable drug delivery systems for the controlled
release of macromolecules, including therapeutic peptides, proteins, and nucleic
acids [7, 11, 12, 126]. However, despite their many favorable characteristics, these
drug delivery systems still have not found their way into broad clinical applications.
One reason is that the quantity and homogeneity of drug loading are still a matter of
concern [11, 12]. Moreover, the high water content of most hydrogels often results
in relatively rapid drug release over a few hours to several days. This not only
shortens the efficacy of the applied drug delivery system, but also carries the risk of
provoking harmful side-effects [7, 11, 12]. Therefore, research focuses on extending the
duration of drug release, and expanding the range of molecules which can be effectively
delivered [11, 12]. Because the resulting release profiles are hardly predictable, the
developed formulations are usually characterized by release studies. The reliability of
these experiments is, however, limited. The results depend on a number of parameters,
such as the geometry of the dosage form, its swelling capacity, and the type and
amount of release medium [127, 128]. Furthermore, release experiments are often
time-consuming, and require relatively large drug loads. Depending on the expected
release kinetics, the sampling time can last up to several days. For these reasons,
screening large libraries of potential drug delivery formulations can be a challenging
task. A wider range of fast and reliable analytical methods that directly asses drug
diffusivity would certainly aid further research and accelerate the development of new
hydrogel-based drug delivery systems.
Fluorescence recovery after photobleaching (FRAP) is a well-established ana-
lytical method that has been widely used to study the translational diffusion of
molecules [129–131]. The basic instrumentation for FRAP experiments consists of an
optical microscope equipped with a light source to bleach arbitrary regions within
the sample. Furthermore, some of the probe molecules must be fluorescently labeled.
After bleaching the probe molecules within the region of interest, the fluorescence
recovery due to diffusion of unbleached molecules is recorded; the diffusion coefficients
can then be calculated from the resulting recovery profiles. Although FRAP experi-
ments have already been used for the characterization of hydrogel-based drug delivery
systems [132–136], the relevance of these data for the expected release profiles has
59
Chapter 4 Hydrogel-based drug delivery systems
yet to be proven. Therefore, systematic comparisons between the measured diffusion
coefficients and the resulting release kinetics will be required until FRAP experiments
and other analytical techniques are generally accepted.
In this study, hydrogels were prepared by step-growth polymerization of branched
poly(ethylene glycol) (PEG) and loaded with fluoresceine isothiocyanate (FITC)
labeled dextrans as macromolecular model drugs. Because of their well defined archi-
tecture [74, 117], these gels can be considered model systems for hydrogel-based drug
delivery formulations. To get an estimate of the expected FITC-dextran diffusivity, the
gels were characterized by mechanical testing and equilibrium swelling studies. Subse-
quently, the translational diffusion coefficients of the incorporated FITC-dextrans were
determined by FRAP. In one sample, FITC-dextran diffusivity was also measured by
pulsed field gradient nuclear magnetic resonance (NMR) spectroscopy, an independent
analytical method to study the translational diffusion of molecules [137, 138]. To
complete the experimental data, the apparent diffusion coefficients of the incorporated
FITC-dextrans were also calculated from release experiments. By comparing these
values, we show the significance of mechanical testing and FRAP experiments for the
characterization of hydrogel-based drug delivery systems. Furthermore, we discuss
the limits of these techniques and comment on the possibility of predicting entire
release profiles from the obtained diffusion coefficients.
4.2 Materials and methods
4.2.1 Materials
Acrylonitrile, N,N’ -dicyclohexylcarbodiimide (DCC), diisopropyl azodicarboxylate
(DIAD), fluoresceine isothiocyanate (FITC) labeled dextrans (FITC-dextrans), and
N-hydroxysuccinimide (HOSu) were purchased from Sigma-Aldrich (Taufkirchen,
Germany). Hexane and phthalimide were obtained from Acros Organics (Geel,
Belgium). Four-armed poly(ethylene glycol) (molecular weight 10 kDa, 4armPEG10k-
OH) was purchased from Nektar Therapeutics (Huntsville, AL). Deuterium oxide
(D2O) was obtained from Deutero GmbH (Kastellaun, Germany). Phosphate buffered
60
4.2 Materials and methods
saline (PBS) was purchased from Invitrogen GmbH (Karlsruhe, Germany). All other
chemicals were of analytical grade and purchased from Merck KGaA (Darmstadt,
Germany). Deionized water was obtained from a Milli-Q water purification system
from Millipore (Schwalbach, Germany).
4.2.2 Synthesis of polymers
Branched PEG-succinimidyl propionates (4armPEG10k-SPA) were synthesized as
previously described [139]. In brief, PEG-propionic acid was obtained through
Michael-type addition reaction of 4armPEG10k-OH onto acrylonitrile and subsequent
hydrolysis under alkaline conditions. The obtained carboxylic acid groups were then
converted to amine-reactive succinimidyl ester groups using HOSu and DCC. The
synthesis of branched PEG-amines (4armPEG10k-NH2) also followed a previously
established procedure [139]. First, phthalimido-PEGs were synthesized by reaction
of 4armPEG10k-OH, phthalimide, triphenylphosphine, and DIAD. The phthalimido
groups were then converted into primary amines by hydrazinolysis.
4.2.3 Rheological characterization of hydrogels
Gelation kinetics and gel strength were studied at 25 ◦C by performing oscillatory
shear experiments using a TA Instruments AR 2000 rheometer (TA Instruments,
Eschborn, Germany) with parallel plate geometry. For the preparation of hydrogels,
defined amounts of 4armPEG10k-NH2 (25 or 50 mg) were dissolved in 1000 µL of
25 mM phosphate buffer, pH 7.0. Immediately before starting the experiment, the
polymer solution was added to an equal amount of 4armPEG10k-SPA (25 or 50 mg).
After vigorous stirring, the precursor solution was poured onto the bottom plate of
the rheometer. The upper plate (20 mm in diameter) was then lowered to a gap size
of 1000 µm, and the measurement was started. The evolution of storage (G′) and loss
moduli (G′′) was recorded as a function of time at 1 Hz oscillatory frequency and a
constant strain of 0.05. A solvent trap was used in order to prevent the evaporation
of water. The cross-over of G′ and G′′ was regarded as the gel point. After 120 min,
the complex shear modulus (G∗) was determined. The gel disks were then removed
61
Chapter 4 Hydrogel-based drug delivery systems
from the rheometer, immersed in 10 mL of PBS, and incubated at 37 ◦C for 24 h. To
measure G∗ in the swollen state, the hydrogel disks were again placed in the center
of the lower plate and the measuring gap was slowly closed until a normal force of
approx. 150 mN was reached. The resulting compression was sufficient to prevent
slippage. The following frequency sweep was conducted at constant strain (0.05) as
a function of frequency (from 0.1 to 10 Hz). Finally, an amplitude sweep (from 0.1
to 10, 000 Pa oscillatory stress) was performed in order to confirm that all previous
measurements were conducted within the linear viscoelastic region of the sample. All
experiments were carried out in triplicate and the results are expressed as means ±standard deviations.
4.2.4 Equilibrium swelling of hydrogels
For the swelling studies, defined amounts of 4armPEG10k-NH2 (25 or 50 mg) were
dissolved in 1000 µL of phosphate buffer (25 mM, pH 7.0) and added to an equal
amount of 4armPEG10k-SPA (25 or 50 mg). After vortexing, 250 µL of the precursor
solution were cast into cylindrical glass molds (7 mm inner diameter) and allowed
to gel for 2 h. The samples were then weighed in air and hexane before and after
swelling for 24 h in 10 mL of PBS using a density determination kit (Mettler-Toledo,
Gießen, Germany). Using Archimedes’ principle, the gel volumes after cross-linking
(Vgc) and after swelling (Vgs) were determined. The volume of the dry polymer (Vp)
was calculated from the mass of the freeze-dried hydrogel and the density of the
polymer in the dry state (taken as the density of PEG, 1.12 g · cm−3). With these
parameters, the polymer fraction of the gel after cross-linking, v2c = Vp/Vgc, and in
the swollen state, v2s = Vp/Vgs, was calculated. The reciprocal of v2s is usually called
the volumetric swelling ratio (Q). All experiments were carried out in triplicate and
the results are expressed as means ± standard deviations.
62
4.2 Materials and methods
4.2.5 Calculation of hydrogel network mesh size
The number of moles of elastically active chains in the hydrogel network, νe, can be
calculated from equilibrium swelling measurements using a modified version of the
classical Flory-Rehner equation [140–142]:
νe = − VpV1v2c
· [ln(1− v2s) + v2s + χ1v22s][(
v2sv2c
) 13 − 2
f
(v2sv2c
)] (4.1)
In this equation, χ1 is the Flory-Huggins interaction parameter for PEG in water
(taken as 0.43), V1 is the molar volume of the solvent (18 cm3 ·mol−1), and f is the
functionality of the cross-links (4 in the case of four-armed PEG). In the case of
perfectly formed condensation gels, the average molecular weight between cross-links,
Mc, can be calculated by Mc = mp/νe, where mp is the total mass of PEG in the
hydrogel [142]. Using this parameter, the average mesh size (ξ) can be calculated
by [143]:
ξ = v− 1
32s l
(2Mc
Mr
) 12
C12n (4.2)
where l is the bond length along the polymer backbone (taken as 0.146 nm), Mr is
the molecular mass of the PEG repeating unit (44 g ·mol−1), and Cn is the Flory
characteristic ratio (here taken as 4 for PEG) [144].
Alternatively, information about the network structure can be derived from the
equilibrium modulus, Ge, which provides an independent estimate of νe. The elastic
behavior of gel networks thereby falls between two idealized limits, affine and phantom,
which relate the network structure to the elastic deformation under applied stress [145].
It has been shown, however, that for gels cross-linked at low concentrations, the
agreement between the phantom model and the experimental observations appears
to be satisfactory [146]. Utilizing the following expression, νe can be obtained from
Ge [145, 146]:
Ge = RT
(νe − µeVgc
)v− 1
32c v
132s (4.3)
63
Chapter 4 Hydrogel-based drug delivery systems
Here, R is the molar gas constant (8.3145 J ·mol−1 ·K−1), T the absolute temperature
(298.16 K), and µe refers to the number of moles of cross-links. For a network with
only one type of cross-link, νe − µe = νe(1− 2/f) [145].
4.2.6 Fluorescence recovery after photobleaching (FRAP)
To investigate the mobility of incorporated macromolecules, the hydrogels were loaded
with different FITC-dextrans (20, 150, and 2, 000 kDa molecular weight). The samples
were prepared by dissolving defined amounts of 4armPEG10k-NH2 (12.5 or 25 mg) in
450 µL of phosphate buffer (25 mM, pH 7.0). Subsequently, 50 µL of the respective
FITC-dextran stock solution (prepared in the same buffer, c = 10 mg/mL) were
added. This mixture was then added to an equal amount of 4armPEG10k-SPA
(12.5 or 25 mg), vigorously stirred, and cast into cylindrical glass molds (5 mm inner
diameter, 5 mm height). After gelling for 2 h, the samples were removed from the glass
molds, placed into a Lab-TekTM
II Chambered Coverglass (Thermo Fisher Scientific,
Langenselbold, Germany), warmed to 37 ◦C, and positioned on the microscope stage.
The FRAP experiments were performed on a Zeiss Axiovert 200 M microscope
coupled to a LSM 510 META scanning device (Carl Zeiss MicroImaging GmbH,
Jena, Germany). A Plan-Neofluar 10 × objective lens with a numerical aperture of
0.30 was used. All bleaching experiments were performed using the 488 nm-line of
a 30 mW Ar-ion laser operating at 25 % output power. The confocal pinhole was
opened completely in order to detect as much fluorescence as possible. After the
location of interest was brought into focus, a time-series of digital images with a
resolution of 512 × 128 pixel was recorded using a highly attenuated laser beam
(0.2 % transmission). The interval between two consecutive images was between 1
and 4 s, depending on the expected diffusion time of the used FITC-dextran. After
the acquisition of five prebleach images, a uniform disk with a diameter of 36 µm
was bleached at maximum laser intensity (100 % transmission). The bleaching phase
usually took 0.8 – 1.6 s, which should be sufficiently short to avoid fluorescence
recovery during bleaching [130, 131]. Immediately after bleaching, a stack of 75
images was acquired at low laser intensity (0.2 % transmission) in order to measure
the recovery of fluorescence inside the bleached area. To extract the experimental
64
4.2 Materials and methods
recovery curve from the image stack, the mean fluorescence intensities inside the
bleached region, Ifrap(t), and inside a reference region, Iref (t), were calculated for each
time point t using the NIH software ImageJ. In the next step, Ifrap(t) was normalized
to the prebleach intensity, Ifrap(pre), and corrected for any bleaching effects that
might have occurred during image acquisition:
f(t) =Iref (pre)
Iref (t)· Ifrap(t)
Ifrap(pre)(4.4)
Here, f(t) is the normalized fluorescence intensity inside the bleached region, and
Iref (pre) is the fluorescence intensity inside the reference region before bleaching. In
the following step, f(t) was further normalized to the full scale using:
F (t) =f(t)− f(0)
f(pre)− f(0)(4.5)
where f(0) is the normalized fluorescence intensity immediately after bleaching,
and f(pre) the normalized fluorescence intensity before bleaching. Finally, the
characteristic diffusion time τD and the mobile fraction k were determined by a
least-squares fit of the following expression to the experimental recovery curve:
F (t) = k · e−τD2t
[I0
(τD2t
)+ I1
(τD2t
)](4.6)
where I0 and I1 are the modified Bessel functions of the first kind of zero and first
order. This FRAP model has been described earlier by Soumpasis for a uniform
disk bleached in two-dimensional samples [147]. However, if an objective lens of low
numerical aperture is used for bleaching, Equation (4.6) can also be used to describe
the fluorescence recovery in three-dimensional samples [130]. The diffusion coefficient
(D) was then calculated by D = w2/τD, where w is the radius of the bleached spot
(18 µm in all experiments).
In order to measure the diffusion coefficients also in the swollen state of the
hydrogels, the samples were immersed in 500 µL of PBS which contained 1 mg/mL of
the respective FITC-dextran. After 24 and 48 h of incubation at 37 ◦C, the swollen
gels were removed, rinsed with PBS, and placed into a Lab-TekTM
II Chambered
Coverglass again. The FRAP experiments were then repeated as described above.
65
Chapter 4 Hydrogel-based drug delivery systems
4.2.7 Nuclear magnetic resonance (NMR) spectroscopy
For the NMR experiments, hydrogels were prepared in deuterated buffer solutions. For
this purpose, 68.05 mg KH2PO4 were dissolved in 20 mL of D2O. This solution was
adjusted with NaOH to a pH meter reading of 7.0 and lyophilized; the lyophilisate was
then reconstituted with 20 mL of D2O. To prepare a hydrogel, 25 mg 4armPEG10k-
NH2 were dissolved in 900 µL of this buffer solution and mixed with 100 µL of
a solution containing 10 mg/mL FITC-dextran (molecular weight 20 kDa). This
mixture was then added to 25 mg 4armPEG10k-SPA, vortexed, transferred into a
NMR tube, and allowed to gel for 2 h.
Diffusion ordered NMR spectroscopy (DOSY) experiments were performed on a
Bruker AVANCE 600 spectrometer (Bruker BioSpin GmbH, Rheinstetten, Germany)
operating at 600.13 MHz. As pulse sequence, a stimulated echo diffusion experiment
with bipolar gradients, longitudinal eddy current delay, and two spoil gradients was
chosen. Preparatory experiments revealed that transversal relaxation was not critical
(data not shown). To minimize the eddy current effects after long gradients, sinusoidal
gradient shapes were selected. The gradient strength was varied linearly from 5 %
to 95 % of the maximal available 53.5 G/cm. To study FITC-dextran diffusivity,
DOSY spectra were acquired at 300 K, and the attenuation of the NMR signal at
4.98 ppm was observed. Typical values of the parameters were 2 s for the relaxation
delay, 150 ms for the diffusion time (∆), and 8.5 ms for the gradient pulse length (δ).
Diffusion coefficients were extracted out of the signal decay curve using the SimFit
algorithm of the Bruker TopSpin Software.
4.2.8 Release of FITC-dextrans
As described for the FRAP experiments, the hydrogels were loaded with different
FITC-dextrans (molecular weight 20, 150, and 2, 000 kDa). To prepare the gel
samples, defined amounts of 4armPEG10k-NH2 (12.5 or 25 mg) were dissolved in
450 µL of phosphate buffer (25 mM, pH 7.0) and mixed with 50 µL of the respective
FITC-dextran stock solution (prepared in the same buffer, c = 10 mg/mL). This
mixture was then added to an equal amount of 4armPEG10k-SPA (12.5 or 25 mg)
and vigorously stirred. In each case, 100 µL of this precursor solution were cast into
66
4.3 Results and discussion
cylindrical glass molds (5 mm inner diameter, 5 mm height) and allowed to gel for
2 h. Afterwards, the gels were removed from the glass molds, immersed in 500 µL of
PBS which contained 1 mg/mL of the respective FITC-dextran, and incubated for
24 h at 37 ◦C. After swelling, the gels were removed, rinsed with PBS and measured
using a slide rule. The gels were placed into glass vials, covered with 10 mL of PBS
(containing 0.025 % sodium azide), and maintained at 37 ◦C in a shaking water bath
(approx. 50 rpm). At specified time intervals, 500 µL of the release medium were
collected and replaced with fresh buffer. The amounts of released FITC-dextrans
(λex = 490 nm, λem = 520 nm) were determined on a PerkinElmer LS 55 Fluorescence
spectrometer (PerkinElmer, Wiesbaden, Germany) using standard calibration curves.
Considering perfect sink conditions throughout the experiment and the fact that
FITC-dextran release is purely diffusion controlled, the following solution of Fick’s sec-
ond law of diffusion can be used to describe the experimental release profiles [128, 148]:
Mt
M∞= 1− 32
π2
∞∑n=1
1
q2n
exp
(−q
2n
r2Dt
)·∞∑p=0
1
(2p+ 1)2exp
(−(2p+ 1)2π2
h2Dt
)(4.7)
where Mt and M∞ represent the absolute cumulative amounts of FITC-dextran
released at time t and infinity, respectively; the qn are the roots of the Bessel function
of the first kind of zero order, and r and h denote the radius and height of the gel
cylinders. By a least squares fit of Equation (4.7) to the experimental release profiles,
the apparent diffusion coefficients (D) of the FITC-dextrans were determined.
4.3 Results and discussion
4.3.1 Physicochemical characterization of hydrogels
Hydrogels were prepared by step-growth polymerization of branched PEG-amines
with branched PEG-succinimidyl propionates. At the beginning of the experiments,
all samples behaved like free-flowing liquids (G′′ > G′). Gelation occurred after
8.1± 0.2 min for gels containing 5 % polymer, and 3.2± 0.1 min for gels containing
10 % polymer (Table 4.1). During the course of the experiment, cross-linking further
67
Chapter 4 Hydrogel-based drug delivery systems
proceeded as indicated by the steadily increasing value of G′. After approx. 120 min,
the value of G′ reached a plateau and exceeded that of G′′ by several orders of
magnitude (G′ � G′′). The gel strength (|G∗|) was 2134 ± 89 Pa for hydrogels
containing 5 % polymer, and 6808±145 Pa for gels containing 10 % polymer (Table 4.1).
After swelling in PBS, G∗ was measured at constant strain as a function of frequency.
Within the studied range, the absolute value of G∗ was almost insensitive to the
oscillatory frequency, which is typical for covalently cross-linked networks [149]. In the
swollen state, the gel strength (determined at a frequency of 1 Hz) was 1873± 183 Pa
for gels containing 5 % polymer, and 5767±149 Pa for those containing 10 % polymer
(Table 4.1). In amplitude sweep measurements, both gel types showed a broad linear
viscoelastic region up to an oscillatory stress of approx. 300 and 750 Pa, respectively,
indicating that all previous experiments were run well within the linear viscoelastic
region of the samples.
Table 4.1: Physicochemical characteristics of the prepared hydrogels
5 % Polymer 10 % Polymer
Gelation time (min) 8.1± 0.2 3.2± 0.1
Gel strength |G∗| after cross-linking (Pa) 2134± 89 6808± 145
Gel strength |G∗| after swelling (Pa) 1873± 183 5767± 149
Volumetric swelling ratio (Q) 32.2± 0.1 24.0± 0.1
Mc (g ·mol−1 ) calculated from swelling data 6652± 70 6547± 57
ξ (nm) calculated from swelling data 16.1± 0.1 14.5± 0.1
Mc (g ·mol−1 ) calculated from Ge 35279 18775
ξ (nm) calculated from Ge 37.2 24.6
In general, the absolute value of G∗ of hydrogels containing 10 % polymer was
approx. three times higher than that of gels containing only 5 % polymer. In a
defect-free, end-linked network, however, the mechanical characteristics should be
independent of the precursor concentration [74]. As this was obviously not the
case, network defects seem to play a significant role in the structure of the prepared
hydrogels. At low precursor concentrations, the concentration of elastically active
chains is probably reduced by non-reacted groups and loops, whereas it is increased
by entanglements at higher precursor concentrations.
68
4.3 Results and discussion
A similar trend was observed in the volumetric swelling ratio. The value of Q
decreased from 32.2±0.1 (5 % polymer) to 24.0±0.1 (10 % polymer). Using a modified
version of the Flory-Rehner equation, Mc was calculated to be 6652 ± 70 g ·mol−1
for gels containing 5 % polymer, and 6547± 57 g ·mol−1 for those containing 10 %
polymer. These values correspond to an average mesh size (ξ) of 16.1 ± 0.1 and
14.5± 0.1 nm, respectively (Table 4.1). Given the large differences in the absolute
values of G∗, however, it seems unlikely that the real values of ξ are close to each
other. The problems associated with the estimation of ξ can be attributed to the
fact that the Flory-Huggins interaction parameter (χ1) has not been experimentally
determined. The value of χ1 (0.43) was originally calculated for linear PEG chains
in water [150] and might differ for the macromers used in this study. Furthermore,
Equation (4.1) does not account for any network defects (such as elastically inactive
dangling ends) as this would require the number of non-reacted groups [151]. For
these reasons, the network parameters were also calculated from the equilibrium
modulus (Ge) of the hydrogels. Since G∗ was almost constant within the studied
frequency range, the absolute value of G∗ determined at 1 Hz oscillatory frequency
was taken as equal to Ge. The resulting values of ξ were 37.2 and 24.6 nm for gels
prepared from 5 % and 10 % polymer, respectively (Table 4.1). These values seem to
be more realistic given the observed experimental data.
4.3.2 Estimation of diffusion coefficients
Three different FITC-dextrans with molecular weights of 20, 150, and 2, 000 kDa were
used as macromolecular model compounds in order to evaluate the mass transport char-
acteristics of the prepared hydrogels. Their hydrodynamic radii, rH , were calculated
according to Braeckmans et al. [130]. Knowing these values, a theoretical estimate
of their diffusion coefficients in water (D0) can be made with the Stokes-Einstein
equation:
D0 =kT
6πηrH(4.8)
69
Chapter 4 Hydrogel-based drug delivery systems
where k is the Boltzmann constant (1.38 · 10−23 J ·K−1), T the absolute temperature
(310.16 K), and η the dynamic viscosity of water (0.001 Pa · s). The results are
summarized in Table 4.2.
Table 4.2: Hydrodynamic radii (rH) and diffusion coefficients in water (D0) of FITC-dextrans. The values of D0 were estimated using Equation (4.8).
FITC-dextran 20 kDa 150 kDa 2,000 kDa
rH (nm) 2.9 8.3 32.8
D0 (µm2/s ) 79.6 27.4 7.0
Based on a free-volume approach proposed by Lustig and Peppas [126, 152], the
diffusivity of incorporated molecules (Dg) can be related to the structure of the
hydrogel network:
Dg
D0
=
(1− rH
ξ
)· exp
(−Y
(v2s
1− v2s
))(4.9)
Here, Y is defined as the ratio of the critical volume required for a translational
movement of the encapsulated molecule and the average free volume per solvent
molecule. For correlation purposes, a good approximation of Y is unity [126]. With
this equation, the FITC-dextran diffusivities were estimated based on the determined
values of ξ (derived from mechanical testing), and the results are presented in
Table 4.3. According to the calculated values, the smaller FITC-dextrans (20 and
150 kDa molecular weight) should be able to move almost freely within the formed
hydrogels (Dg ∼ D0). For the larger FITC-dextran (2, 000 kDa molecular weight),
however, Equation (4.9) predicts strongly restricted diffusivities. In case of gels
prepared from 10 % polymer, the average network mesh size was lower than the
hydrodynamic radius of the FITC-dextran (ξ < rH) and Dg could not be calculated.
4.3.3 Determination of diffusion coefficients by FRAP
After these theoretical considerations, the diffusion coefficients (D) of the incorpo-
rated FITC-dextrans were measured. A typical FRAP experiment is illustrated in
Figure 4.1A. After acquisition of the image stacks, the mobile fractions k and the
70
4.3 Results and discussion
diffusion coefficients (D) were determined by least-squares fits of Equation (4.6) to the
experimental recovery profiles (see Figure 4.1B). The coefficients of determination were
generally very high (R2 > 0.99), except for gels loaded with 2, 000 kDa FITC-dextrans
(10 % polymer), where slightly lower values were obtained (R2 > 0.89).
A) B)
0.0
0.2
0.4
0.6
0.8
1.0
0 20 40 60 80 100 120 140 160
Time (s)
F(t
)
k = 0.89
D = 5.30 µm2/s
R2
= 0.9985
Figure 4.1: (A) Image stack of a typical FRAP experiment (5 % polymer, FITC-dextran150 kDa). The first image shows the sample before bleaching (prebleach image). Im-mediately before taking the second image, a uniform disk is bleached. Then, the laserintensity is reduced again, and fluorescence recovery is recorded over multiple images.(B) An image processing program extracts the recovery curve from the image stack.The experimental data are indicated by symbols (u). A least-squares fit (solid line) ofEquation (4.6) yields the mobile fraction k and the diffusion coefficient D.
Remarkably, the values of k did not vary significantly over the course of the
experiment. In case of the 20 kDa FITC-dextrans, all molecules were mobile, regardless
of the polymer concentration (see Table 4.3). This was expected, as the hydrodynamic
diameter of this molecule was well below the average network mesh size of the hydrogels.
In contrast, the mobility of the 150 kDa FITC-dextrans was restricted to some extent
(k = 0.89 and k = 0.66 for gels containing 5 % and 10 % polymer, respectively). This
effect was even more pronounced in hydrogels loaded with 2, 000 kDa FITC-dextrans
(k = 0.20 and k = 0.24 for 5 % and 10 % polymer, respectively); this can also be
explained by the size of these molecules. Their hydrodynamic diameters exceeded the
average hydrogel mesh size by approximately a factor of two (Table 4.1 and 4.2). The
calculated diffusion coefficients D (after swelling for 48 h) are presented in Table 4.3.
For all studied FITC-dextrans, the diffusion coefficients were generally slightly
lower in hydrogels containing 10 % polymer than in gels prepared from only 5 %
polymer. As shown in Figure 4.2, an increase in D during hydrogel swelling was
71
Chapter 4 Hydrogel-based drug delivery systems
Table 4.3: Measurement of the mobile fraction k and diffusion coefficient D (µm2/s ) ofFITC-dextrans incorporated into hydrogels. Data are presented as means ± standarddeviations (n = 3 for each group). For comparison, the expected diffusion coefficients Dg
(µm2/s ) are also included.
FITC-dextran 20 kDa 150 kDa 2,000 kDa
Polymer conc. 5 % 10 % 5 % 10 % 5 % 10 %
Dg (expected)a 71.1 67.3 20.6 17.3 0.8 –
k (FRAP)b,c 1.04 1.03 0.89 0.66 0.20 0.24
D (FRAP)c 20.2 ± 0.2 18.1 ± 0.1 15.2 ± 0.1 12.8 ± 0.1 0.9 ± 0.1 0.1 ± 0.0
D (DOSY) 30.4 – – – – –
D (release) 47.7 ± 1.1 44.7 ± 1.2 24.5 ± 0.6 25.8 ± 0.7 3.7 ± 0.7 2.2 ± 0.7
aCalculated from the average network mesh size using Equation (4.9)bData are presented as means; standard deviations (values < 0.03) are not shown for claritycDetermined after swelling in PBS for 48 h
observed for all smaller FITC-dextrans (20 and 150 kDa molecular weight). After
approx. 48 h, however, the diffusion coefficients had levelled off to their final values. As
expected, the diffusion coefficients of the largest FITC-dextran (2, 000 kDa molecular
weight) were very low (0.9 ± 0.1 and 0.1 ± 0.0 µm2/s for gels containing 5 % and
10 % polymer, respectively). In case of the 150 kDa FITC-dextrans, the measured
values of D also correlated well with the predicted values (15.2± 0.1 vs. 20.6 µm2/s
for gels containing 5 % polymer, and 12.8± 0.1 vs. 17.3 µm2/s for hydrogels prepared
from 10 % polymer, Table 4.3). However, the FRAP technique yielded lower values
for the diffusion coefficients of the 20 kDa FITC-dextrans than expected (20.2± 0.2
and 18.1 ± 0.2 µm2/s for gels prepared from 5 % and 10 % polymer, respectively).
Since these molecules were expected to diffuse very fast in the gels (Dg = 71.1 and
67.3 µm2/s, Table 4.3), significant recovery of fluorescence might have occurred during
the bleaching phase, which would consequently lead to an underestimation of D.
4.3.4 Determination of diffusion coefficients by NMR
To get more accurate values of D for the smallest FITC-dextran (20 kDa molecular
weight), the diffusivity of this macromolecule was examined by pulsed field gradient
72
4.3 Results and discussion
0
5
10
15
20
25
FITC-dextran 20 kDa FITC-dextran 150 kDa FITC-dextran 2000 kDa
Dif
fusio
nco
eff
icie
nt
(µm
²/s)
5 % polymer, non-swollen (t = 0 h) 10 % polymer, non-swollen (t = 0 h)
5 % polymer, swollen (t = 24 h) 10 % polymer, swollen (t = 24 h)
5 % polymer, swollen (t = 48 h) 10 % polymer, swollen (t = 48 h)
Figure 4.2: Diffusion coefficients (D) of FITC-dextrans determined by FRAP. Data areshown as means ± standard deviations (n = 3).
NMR spectroscopy. Using a magnetic field gradient, the probe molecules get spatially
labeled according to their position in the NMR tube. If diffusion occurs, the NMR
signal intensity is attenuated depending on the diffusion time (∆), the length of
the gradient (δ), and the gradient strength. In two-dimensional DOSY experiments,
usually the strength of the magnetic field gradient is varied, whereas ∆ and δ are
kept constant. In the resulting spectra, the chemical shifts are plotted along the F2
axis and the diffusion coefficients along the F1 axis.
For the studied FITC-dextran, a diffusion coefficient of 30.38 µm2/s was extracted
from the signal decay curve. Although non-swollen gel samples were used for NMR
measurements, the calculated value of D was considerably higher than that determined
by FRAP after 48 h of swelling (Table 4.3). Therefore, it can be expected that the
measured values of D have good chances to agree with the predicted ones, if NMR
measurements could be performed in swollen hydrogel samples. Altogether, pulsed
field gradient NMR spectroscopy is regarded as a valuable and robust method to
determine the diffusion coefficients of comparatively fast-diffusing molecules. In
addition, it is particularly advantageous as non-fluorescent probe molecules can also
73
Chapter 4 Hydrogel-based drug delivery systems
be studied. In the case of slow-diffusing macromolecules, however, DOSY experiments
are less well suited, as the signal intensity will not decay sufficiently within acceptable
limits of ∆ and δ. For this reason, the diffusion coefficients of the two larger FITC-
dextrans (150 and 2, 000 kDa molecular weight) could not be determined by pulsed
field gradient NMR spectroscopy.
4.3.5 Release of FITC-dextrans
In the final experiment, the in vitro FITC-dextran release was studied. For this
purpose, FITC-dextran loaded gel cylinders were prepared and incubated in a com-
paratively large amount of PBS in order to assure sink conditions. To calculate
the diffusion coefficients of the incorporated FITC-dextrans, an analytical solution
of Fick’s second law of diffusion [Equation (4.7)] was fitted to the experimentally
determined release kinetics. Since this theory does not account for hydrogel swelling
(diameter and height of the gel cylinder are assumed to be constant), all samples
were pre-swollen in PBS which contained 1 mg/mL of the respective FITC-dextran.
As a consequence of this, the exact amount of FITC-dextran loading was unknown
(actual FITC-dextran load per hydrogel ≥ 100 µg), and only the absolute values of
the released quantities were represented.
As can be seen in Figure 4.3, good agreement between theoretical values (solid lines)
and experimental data (symbols) was obtained in all cases (R2 > 0.98). The apparent
diffusion coefficients of the investigated FITC-dextrans (20, 150, and 2, 000 kDa molec-
ular weight) are listed in Table 4.3. As already observed in the FRAP experiments,
the differences between the two polymer concentrations (5 % and 10 %) were generally
very small. In all cases, the determined values of D were also significantly higher
than the values measured by FRAP. Such discrepancies have been described in the
literature and are most likely due to the experimental setup [132, 135, 136]. In release
experiments, mass transport occurs along concentration gradients (classical Fickian
diffusion), whereas FRAP experiments investigate processes of self-diffusion (diffusion
in the absence of concentration gradients). Furthermore, the larger FITC-dextrans
(2, 000 kDa molecular weight) in particular show a relatively broad molecular weight
distribution [153]. Therefore it might be possible that two different populations of
74
4.4 Conclusion
0
20
40
60
80
100
120
140
160
0 24 48 72 96 120 144 168 192
Time (h)
Ab
so
lute
rele
ased
am
ou
nt
(µg
)
5 % polymer, FITC-dextran 20 kDa 10 % polymer, FITC-dextran 20 kDa
5 % polymer, FITC-dextran 150 kDa 10 % polymer, FITC-dextran 150 kDa
5 % polymer, FITC-dextran 2000 kDa 10 % polymer, FITC-dextran 2000 kDa
Figure 4.3: Absolute cumulative amounts of released FITC-dextrans. Data are shown asmeans ± standard deviations (n = 3). Symbols represent the experimental data, solidlines the theoretical values obtained from Equation (4.7).
FITC-dextrans are present within the hydrogels: a low molecular weight fraction
with D almost equal to that in water, and a high molecular weight fraction with
restricted mobility. As outlined in section 4.3.3, the fraction of low molecular weight
FITC-dextrans probably diffuses too fast in order to determine D using FRAP. Con-
sequently, only molecules with restricted mobility will account for the recovery profile,
which results in lower values of D.
4.4 Conclusion
Hydrogels were prepared by step-growth polymerization of branched PEG-amines
with branched PEG-succinimidyl propionates and loaded with three different FITC-
dextrans (20, 150, and 2, 000 kDa molecular weight) as macromolecular model drugs.
Mechanical tests and swelling studies allowed prediction of the FITC-dextran mobility.
The actual diffusion coefficients D inside the gels were measured by fluorescence
75
Chapter 4 Hydrogel-based drug delivery systems
recovery after photobleaching (FRAP) and pulsed field gradient NMR spectroscopy.
In a final experiment, the apparent diffusion coefficients of incorporated FITC-
dextrans were determined by least-squares fits of appropriate model equations to
experimentally obtained release profiles. FRAP proved to be suitable to measure
the diffusion coefficients of slow-diffusing molecules, whereas pulsed field gradient
NMR spectroscopy appears to be especially suited to determine D of fast-diffusing
substances. Furthermore, FRAP could be used to follow changes in drug diffusivity
during hydrogel swelling and degradation or to measure local diffusion coefficients
in inhomogeneous samples. Altogether, mechanical testing, FRAP, and pulsed field
gradient NMR spectroscopy proved to be valuable tools for the characterization
hydrogel-based drug delivery systems. Although the determined values of D did not
match the apparent diffusion coefficients exactly, the relative differences between
the two polymer concentrations and the different FITC-dextrans were represented
correctly. Measuring D will, therefore, allow evaluation of the potential of newly
developed drug delivery systems within a few minutes. This is a clear advantage over
traditional release experiments which often span a period up to several days. Provided
that appropriate mathematical models exist and the mechanisms controlling drug
release are known, entire release profiles could be predicted. This might contribute to
a better understanding of hydrogel-based materials and accelerate the development
of marketable drug delivery systems.
76
Chapter 5
Biodegradable hydrogels fortime-controlled release of tetheredpeptides or proteins
Ferdinand Brandl1, Jorg Teßmar1, Torsten Blunk1, Achim Gopferich1
1 Department of Pharmaceutical Technology, University of Regensburg, 93040 Regensburg
Submitted to Biomacromolecules.
77
Chapter 5 Biodegradable hydrogels for time-controlled release
Abstract
Tethering drug substances to a gel network is an effective way of controlling the
release kinetics of hydrogel-based drug delivery systems. Here, we report on in situ
forming, biodegradable hydrogels that allow for the covalent attachment of peptides
or proteins. Hydrogels were prepared by step-growth polymerization of branched
poly(ethylene glycol). The gel strength ranged from 1075 to 2435 Pa; the degradation
time varied between 24 and 120 h. Fluorescence recovery after photobleaching showed
that fluorescently labeled bovine serum albumin (FITC-BSA) was successfully bound
to the gel network during gel formation. Within 168 h, the mobility of the tethered
molecules gradually increased due to polymer degradation. Using FITC-BSA and
lysozyme as model proteins, we showed the potential of the developed hydrogels for
time-controlled release. The obtained release profiles had a sigmoidal shape and
matched the degradation profile very well; protein release was complete after 96 h.
78
5.1 Introduction
5.1 Introduction
Over the past decades, hydrogels have been extensively used as matrices for controlled
drug delivery and tissue engineering applications [7, 10–12, 102, 154]. The popularity
of these materials is based on their enormous chemical versatility, which allows
for the design of a broad range of hydrogels with varying properties [10–12, 154].
Furthermore, hydrogel-based materials generally show excellent biocompatibility
because of their physicochemical similarity to the native extracellular matrix [10–12,
154]. Finally, compared to other commonly used biomaterials, such as poly(lactic acid)
or poly(lactide-co-glycolide), hydrophilic polymers are well suited for the entrapment
of biomolecules due to their lack of hydrophobic interactions, which can affect the
activity of these fragile species [11, 154].
Despite this multitude of advantageous characteristics, hydrogels also have several
limitations. As a result of their high water content, most hydrogels usually restrict
the mobility of encapsulated peptides or proteins only moderately. Consequently,
a large amount of the incorporated molecules will be released within a few hours,
which illustrates the necessity for more sophisticated strategies in order to prolong
drug release [7, 11, 12]. Adjusting the cross-linking density is a common approach to
reduce gel permeability and slow down the release kinetics. However, this approach is
only effective in controlling the release of relatively large molecules, such as proteins
with molecular weights of several thousand Daltons. Furthermore, increasing the
cross-linking density often results in decreased hydrophilicity and hence decreased
biocompatibility [11]. Tethering drug substances to the gel network via degradable
anchor groups would be a further strategy to effectively control drug release. In
these systems, the release kinetics is ideally controlled by the degradation of the
anchor group while drug diffusivity only plays a secondary role [7, 11, 12, 155]. How-
ever, the drug molecules often need to be chemically modified in order to allow
for their covalent attachment, which makes these drug delivery systems highly com-
plex [156–158]. Furthermore, drug conjugation may also decrease bioactivity, especially
when peptides or proteins are the target therapeutics [11]. In addition, the anchor
groups have to be carefully designed in order to avoid unwanted side-effects during
degradation in vivo [11].
79
Chapter 5 Biodegradable hydrogels for time-controlled release
N
O
O
OH
NH2
OO
ONH
O
O NH
O
OO
+
ONH
O
O O
O
N
O
O
ONH
O
O NH
O
CO2
ONH
O
OH NH2
Figure 5.1: Schematic illustration of hydrolytically degradable hydrogels for time-controlledrelease of tethered peptides or proteins. For gel formation, branched PEG-succinimidylcarbonates are reacted with branched PEG-amines (lower inset). The same chemistryallows for the immobilization of dissolved peptides or proteins via pendant PEG chains.During gel degradation, these anchoring PEG-chains are cleaved, and the unmodifiedpeptide or protein will be released into solution (upper inset).
In this paper, we report the preparation and characterization of in situ cross-
linkable, biodegradable hydrogels for time-controlled release of peptides or proteins.
For this purpose, branched poly(ethylene glycol) (PEG) was functionalized with
aromatic succinimidyl carbonate groups that readily react with amino groups of other
polymers, peptides, or proteins under formation of biodegradable carbamate bonds.
Aromatic succinimidyl carbonates have been originally described as linking groups
for the reversible PEGylation of proteins [159, 160]. While extensively applied for
the preparation of soluble protein prodrugs, this chemistry has never been used for
the formation of hydrogels. For the preparation of covalently cross-linked hydrogels,
PEG-succinimidyl carbonates were reacted with branched PEG-amines (Figure 5.1,
lower inset). Since the chemical environment of the reacting amino groups was
expected to influence gel formation and degradation, the PEG-amines were further
80
5.2 Materials and methods
modified with alanine, 6-aminohexanoic acid, and lysine. These amino acid moieties
were chosen to represent the different amino groups available in proteins (amino
terminus and ε-amino groups of lysine residues). During gelation, dissolved peptides
or proteins are expected to be immobilized within the gel network via pendant PEG
chains (Figure 5.1). This will improve handling and flexibility of the drug delivery
system, since any peptide or protein could be incorporated without requiring chemical
modifications of these molecules. Similar to soluble prodrugs, the anchoring PEG
chains are cleaved during gel degradation, and the native peptide or protein will be
released into solution (Figure 5.1, upper inset) [159–162]. Non-degradable gels served
as a control group. The prepared hydrogels were characterized with respect to their
mechanical properties, swelling capacities, and degradation rates. To prove their
suitability as potential drug delivery systems, the gels were loaded with fluoresceine
isothiocyanate (FITC) labeled bovine serum albumin (BSA) and lysozyme as model
proteins. The successful immobilization and subsequent liberation of FITC-BSA was
investigated by fluorescence recovery after photobleaching (FRAP) experiments. And
finally, the in vitro release kinetics of FITC-BSA and lysozyme were determined.
5.2 Materials and methods
5.2.1 Materials
Hexane, 3-(4-hydroxyphenyl)propionic acic, methylene chloride (DCM), and phthal-
imide were purchased from Acros Organics (Geel, Belgium). Boc-protected alanine
(Boc-Ala-OH), Boc-6-aminohexanoic acid, and di-Boc protected lysine dicyclohexylam-
monium salt (Boc-Lys(Boc)-OH ·DCHA) were obtained from Bachem GmbH (Weil am
Rhein, Germany). Deuterated chloroform (CDCl3) was obtained from Deutero GmbH
(Kastellaun, Germany). Phosphate buffered saline (PBS) was purchased from Invitro-
gen GmbH (Karlsruhe, Germany). Four-armed poly(ethylene glycol) (molecular weight
10 kDa, 4armPEG10k-OH) was purchased from Nektar Therapeutics (Huntsville, AL).
Acetonitrile, Coomassie Brilliant Blue G250, N,N’ -dicyclohexylcarbodiimide (DCC),
fluoresceine isothiocyanate (FITC) labeled bovine serum albumin (BSA), FITC-
81
Chapter 5 Biodegradable hydrogels for time-controlled release
dextran (150 kDa molecular weight), N-hydroxysuccinimide (HOSu), diisopropyl
azodicarboxylate, N,N’ -diisopropylethylamine (DIPEA), lysozyme (from chicken egg
white), Sigmacote R©, and N,N’ -discuccinimidyl carbonate (DSC) were obtained from
Sigma-Aldrich (Taufkirchen, Germany). Ethanol, ethyl acetate, and diethyl ether
were of technical grade and used without further purification. All other chemicals
were of analytical grade and purchased from Merck KGaA (Darmstadt, Germany).
Deionized water was obtained using a Milli-Q water purification system from Millipore
(Schwalbach, Germany).
5.2.2 Synthesis of PEG-amines
Branched PEG-amines (10 kDa molecular weight, 4armPEG10k-NH2) were synthesized
as previously described [139]. First, phthalimido-PEGs were obtained by reaction of
4armPEG10k-OH, phthalimide, triphenylphosphine, and diisopropyl azodicarboxylate.
The phthalimido groups were then converted into primary amines by hydrazinolysis.
5.2.3 Synthesis of non-degradable PEG-succinimidyl carbonates(4armPEG10k-SC)
2.0 g of 4armPEG10k-OH (0.2 mmol) and 0.82 g of DSC (3.2 mmol) were dissolved
in 20 mL of dried acetonitrile. Then, 520 µL of pyridine (6.4 mmol) were added, and
the reaction mixture was stirred overnight at room temperature. The next day, the
solvent was evaporated under reduced pressure, and 20 mL of DCM were added to
the residue. The insoluble residue was filtered off, and the solution was concentrated
under reduced pressure. The product was crystallized at 0 ◦C under vigorous stirring
by drop-wise addition of diethyl ether. For further purification, the crystallization
step was repeated. The precipitate was collected by filtration, washed with cold
diethyl ether, and dried under vacuum to yield 1.9 g (90 %) of 4armPEG10k-SC
(Figure 5.2A).1H-NMR (4armPEG10k-SC, DMSO-d6, 600 MHz): δ 2.80 ppm (s, 16H, –C(O)CH2-
CH2C(O)–), 3.30 ppm (s, 8H, R3CCH2O–), 3.50 ppm (s, –OCH2CH2O–), 3.69 ppm
(t, 8H, –OCH2CH2OC(O)O–), 4.39 ppm (t, 8H, –OCH2CH2OC(O)O–)
82
5.2 Materials and methods
OO
ONH
O
O
O
ON
O
O
OO
OO
O
ON
O
O
OO
ONH2
OO
ONH
ONH2
CH3
OO
ONH
ONH2
OO
ONH
ONH2
NH2
n
n
n
n
n
n
A)
4armPEG10k-SC (non-degradable)
4armPEG10k-dTyr-SC (degradable)
B)
4armPEG10k-NH2
4armPEG10k-Ala
4armPEG10k-dLys
4armPEG10k-Lys
Figure 5.2: Polymers derived from branched poly(ethylene glycol). Amine-reactive PEG-succinimidyl carbonates (A) and amino acid-modified PEG-amines (B).
5.2.4 Synthesis of degradable PEG-succinimidyl carbonates(4armPEG10k-dTyr-SC)
In the first step, 0.67 g of 3-(4-hydroxyphenyl)propionic acid (4.0 mmol), 0.46 g
of HOSu (4.0 mmol), and 0.83 g of DCC (4.0 mmol) were dissolved in 20 mL of
dried 1,4-dioxane and stirred overnight at room temperature. The next day, the
reaction mixture was passed through a glass filter funnel to remove precipitated
N,N’ -dicyclohexylurea (DCU), combined with a solution of 4armPEG10k-NH2 (5.0 g,
0.5 mmol) and NaHCO3 (0.19 g, 2.2 mmol) in water (20 mL), and stirred overnight
at 50 ◦C. Afterwards, the solvent was evaporated and the residue was dissolved
in water. The raw product was then extracted with DCM. The combined organic
phases were dried over anhydrous Na2SO4, and the solution was concentrated under
reduced pressure. The intermediate was crystallized at 0 ◦C under vigorous stirring
by drop-wise addition of diethyl ether. The precipitate was collected by filtration,
washed with cold diethyl ether, and dried under vacuum to yield 5.1 g (96 %) of
4armPEG10k-dTyr. Thin layer chromatography (TLC) indicated that the product
was free of 4armPEG10k-NH2, 3-(4-hydroxyphenyl)propionic acid, and HOSu.
83
Chapter 5 Biodegradable hydrogels for time-controlled release
1H-NMR (4armPEG10k-dTyr, CDCl3, 600 MHz): δ 2.43 ppm (t, 8H, –C(O)CH2-
CH2–), 2.87 ppm (t, 8H, –C(O)CH2CH2–), 3.41 ppm (s, 8H, R3CCH2O–), 3.64 ppm
(s, –OCH2CH2O–), 6.13 ppm (t, 4H, –NH C(O)–), 6.76 ppm (d, 8H, –C6H4OH),
7.03 ppm (d, 8H, –C6H4OH )
The phenolic hydroxyl groups were then converted into amine-reactive succinimidyl
carbonate groups. For a typical synthesis, 5.3 g of 4armPEG10k-dTyr (0.5 mmol)
and 2.05 g of DSC (8.0 mmol) were dissolved in 50 mL of dried acetonitrile. Then,
1300 µL of pyridine (16.0 mmol) were added, and the reaction mixture was stirred
overnight at room temperature. The next day, the solvent was evaporated under
reduced pressure, and 50 mL of DCM were added to the residue. The insoluble
residue was filtered off, and the solution was concentrated under reduced pressure.
The product was crystallized at 0 ◦C under vigorous stirring by drop-wise addition
of diethyl ether. For further purification, the crystallization step was repeated. The
precipitate was collected by filtration, washed with cold diethyl ether, and dried under
vacuum to yield 5.2 g (93 %) of 4armPEG10k-dTyr-SC (Figure 5.2A).1H-NMR (4armPEG10k-dTyr-SC, CDCl3, 600 MHz): δ 2.48 ppm (t, 8H, –C(O)CH2-
CH2–), 2.88 ppm (s, 16H, –C(O)CH2CH2C(O)–), 2.98 ppm (t, 8H, –C(O)CH2CH2–),
3.41 ppm (s, 8H, R3CCH2O–), 3.64 ppm (s, –OCH2CH2O–), 6.36 ppm (t, 4H,
–NH C(O)–), 7.17 ppm (d, 8H, –C6H4OSu), 7.26 ppm (d, 8H, –C6H4OSu)
5.2.5 Synthesis of alanine-modified PEG-amines(4armPEG10k-Ala)
In the coupling step, 1.51 g of Boc-Ala-OH (8.0 mmol) and 0.83 g of DCC (4.0 mmol)
were dissolved in 20 mL of dried DCM. After stirring for 30 min at room temperature,
the solution was passed through a glass filter funnel to remove precipitated DCU,
and combined with a solution of 4armPEG10k-NH2 (5.0 g, 0.5 mmol) in dried DCM
(50 mL). Then, 700 µL of DIPEA (4.0 mmol) were added, and the mixture was
stirred for 1.5 h at room temperature. The solution was concentrated under reduced
pressure and placed into an ice bath. The intermediate was crystallized at 0 ◦C under
vigorous stirring by drop-wise addition of diethyl ether. The precipitate was collected
84
5.2 Materials and methods
by filtration, washed with cold diethyl ether, and dried under vacuum to yield 5.1 g
(97 %) of 4armPEG10k-Ala-Boc.1H-NMR (4armPEG10k-Ala-Boc, CDCl3, 300 MHz): δ 1.36 ppm (d, 12H, –CH(NH-
Boc)CH3), 1.44 ppm (s, 36H, –OtBu), 3.41 ppm (s, 8H, R3CCH2O–), 3.64 ppm
(s, –OCH2CH2O–), 4.15 ppm (4H, –CH (NHBoc)CH3), 5.20 ppm (4H, –CH(NH Boc)-
CH3), 6.66 ppm (4H, –NH C(O)–)
To deprotect the amino acid moiety, 5.1 g of 4armPEG10k-Ala-Boc (0.5 mmol)
were dissolved in 55 mL of methanolic HCl (prepared by dropping 5 mL of acetyl
chloride into 50 mL of ice-cooled methanol). After stirring for 30 min at room
temperature, the solvent was evaporated, and the residue was dissolved in 25 mL of
water. The raw product was extracted with DCM. The combined organic phases
were dried over anhydrous Na2SO4, and the solution was concentrated under reduced
pressure. 4armPEG10k-Ala was crystallized at 0 ◦C under vigorous stirring by drop-
wise addition of diethyl ether. The precipitate was collected, washed with cold diethyl
ether, and dried under vacuum to yield 4.7 g (96 %) of the product (Figure 5.2B).1H-NMR (4armPEG10k-Ala, CDCl3, 600 MHz): δ 1.34 ppm (d, 12H, –CH(NH2)-
CH3), 3.41 ppm (s, 8H, R3CCH2O–), 3.64 ppm (s, –OCH2CH2O–), 7.55 ppm (t, 4H,
–NH C(O)–)
5.2.6 Synthesis of 6-aminohexanoic acid-modified PEG-amines(4armPEG10k-dLys)
4armPEG10k-dLys (Figure 5.2B) was synthesized from 4armPEG10k-NH2 and Boc-
6-aminohexanoic acid with a 95 % yield as described for 4armPEG10k-Ala.1H-NMR (4armPEG10k-dLys, CDCl3, 600 MHz): δ 1.36 ppm (m, 8H, –CH2CH2-
CH2CH2CH2NH2), 1.47 ppm (m, 8H, –CH2CH2CH2CH2CH2NH2), 1.66 ppm (m, 8H,
–CH2CH2CH2CH2CH2NH2), 2.19 ppm (t, 8H, –CH2CH2CH2CH2CH2NH2), 2.70 ppm
(t, 8H, –CH2CH2CH2CH2CH2NH2), 3.41 ppm (s, 8H, R3CCH2O–), 3.64 ppm
(s, –OCH2CH2O–), 6.23 ppm (t, 4H, –NH C(O)–)
85
Chapter 5 Biodegradable hydrogels for time-controlled release
5.2.7 Synthesis of lysine-modified PEG-amines(4armPEG10k-Lys)
5.0 g of Boc-Lys(Boc)-OH ·DCHA (9.5 mmol) were suspended in 50 mL of ethyl
acetate. A solution of 10 % citric acid in water was added until a clear mixture was
obtained. After stirring for 30 min at 0 ◦C, the aqueous phase was separated and
extracted with ethyl acetate. The combined organic phases were washed with water
until neutral and dried over anhydrous Na2SO4. After evaporation of the solvent,
Boc-Lys(Boc)-OH was dried under vacuum to yield 3.2 g (97 %). 4armPEG10k-Lys
was prepared from 4armPEG10k-NH2 and Boc-Lys(Boc)-OH in a similar manner to
that of 4armPEG10k-Ala with a 95 % yield (Figure 5.2B).1H-NMR (4armPEG10k-Lys, CDCl3, 600 MHz): δ 1.32 – 1.54 ppm (m, 20H,
–CH2CH2CH2CH2NH2), 1.76 – 1.85 ppm (m, 4H, –CH2CH2CH2CH2NH2), 2.69 ppm
(t, 8H, –CH2CH2CH2CH2NH2), 3.32 ppm (4H, –NHC(O)CH (NH2)–), 3.39 ppm (s,
8H, R3CCH2O–), 3.61 ppm (s, –OCH2CH2O–), 7.54 ppm (4H, –OCH2CH2NH C(O)–)
5.2.8 Preparation and rheological characterization of hydrogels
Gelation kinetics and gel strength were studied by performing oscillatory shear
experiments on a TA Instruments AR 2000 rheometer (TA Instruments, Eschborn,
Germany) with parallel plate geometry. For the preparation of hydrogels, accurately
weighed amounts of the amine component were dissolved in 1000 µL of 25 mM
phosphate buffer (Table 5.1) and cooled to 5 ◦C. Immediately before starting the
experiment, this polymer solution was added to the PEG-succinimidyl carbonate
(Table 5.1). The stoichiometric ratio between succinimidyl carbonate and amino
groups was balanced, and the overall polymer concentration was 5 % (w/v) for all
hydrogels. After vortexing, the precursor solution was poured onto the bottom plate
of the rheometer which was also cooled to 5 ◦C. The upper plate (20 mm in diameter)
was then lowered to a gap size of 1000 µm, the temperature was raised to 25 ◦C, and
the measurement was started. The evolution of storage (G′) and loss moduli (G′′)
was recorded as a function of time at 1 Hz oscillatory frequency and a constant strain
of 0.05. A solvent trap was used in order to prevent the evaporation of water. The
cross-over of G′ and G′′ was regarded as the gel point. After 90 min, the absolute
86
5.2 Materials and methods
value of the complex shear modulus (G∗ = G′ + i ·G′′) was determined as a measure
for the gel strength. All experiments were carried out in triplicate and the results are
expressed as means ± standard deviations.
Table 5.1: Composition of the prepared hydrogels. The amine component was dissolved in1000 µL of 25 mM phosphate buffer and added to the PEG-succinimidyl carbonate. Thestoichiometric ratio between succinimidyl carbonate and amino groups was balanced.
GroupPolymer
conc. pH Amine componentPEG-succinimidyl
carbonate
PEG-dLys 5 % 8.0 4armPEG10k-dLys (25 mg) 4armPEG10k-dTyr-SC (25 mg)
PEG-NH2 5 % 7.4 4armPEG10k-NH2 (25 mg) 4armPEG10k-dTyr-SC (25 mg)
PEG-Ala 5 % 6.8 4armPEG10k-Ala (25 mg) 4armPEG10k-dTyr-SC (25 mg)
PEG-Lys 5 % 6.4 4armPEG10k-Lys (17 mg) 4armPEG10k-dTyr-SC (33 mg)
Control 5 % 7.0 4armPEG10k-NH2 (25 mg) 4armPEG10k-SC (25 mg)
5.2.9 Equilibrium swelling of hydrogels and determination ofnetwork parameters
For the swelling studies, accurately weighed amounts of the amine component were
dissolved in 1000 µL of 25 mM phosphate buffer, and added to the PEG-succinimidyl
carbonate (Table 5.1). The stoichiometric ratio between succinimidyl carbonate
and amino groups was balanced, and the overall polymer concentration was again
5 % (w/v) for all gels. After vortexing, 250 µL of the precursor solution were cast
into cylindrical glass molds (7 mm inner diameter) and allowed to gel for 2 h. The
samples were then weighed in air and hexane before and after swelling for 24 h in
10 mL of PBS using a density determination kit (Mettler-Toledo, Gießen, Germany).
To minimize hydrogel degradation, the samples were incubated at 5 ◦C. Based on
Archimedes’ principle, the gel volumes after cross-linking (Vgc) and after swelling (Vgs)
were determined. The volume of the dry polymer (Vp) was calculated from the mass
of the freeze-dried hydrogel and the density of the polymer in the dry state (taken
as the density of PEG, 1.12 g · cm−3). With these parameters, the polymer fraction
of the gel after cross-linking, v2c = Vp/Vgc, and in the swollen state, v2s = Vp/Vgs,
87
Chapter 5 Biodegradable hydrogels for time-controlled release
was calculated. The reciprocal of v2s is usually termed as the volumetric swelling
ratio (Q).
The number of moles of elastically active chains in the hydrogel network, νe, was
calculated using a modified version of the classical Flory-Rehner equation [140–142]:
νe = − VpV1v2c
· [ln(1− v2s) + v2s + χ1v22s][(
v2sv2c
) 13 − 2
f
(v2sv2c
)] (5.1)
Here, χ1 is the Flory-Huggins interaction parameter for PEG in water (taken as 0.43),
V1 is the molar volume of the solvent (18 cm3 ·mol−1), and f is the functionality of
the cross-links (4 in the case of four-armed PEG). Assuming a defect-free network,
the average molecular weight between cross-links, Mc, was calculated by Mc = mp/νe,
where mp is the total mass of PEG in the hydrogel. With this parameter, the average
network mesh size (ξ) was estimated by [143]:
ξ = v− 1
32s l
(2Mc
Mr
) 12
C12n (5.2)
where l is the bond length along the polymer backbone (taken as 0.146 nm), Mr is
the molecular mass of the PEG repeating unit (44 g ·mol−1), and Cn is the Flory
characteristic ratio (here taken as 4 for PEG) [144]. All experiments were carried out
in triplicate and the results are expressed as means ± standard deviations.
5.2.10 Degradation of hydrogels
The gel samples were prepared as described in section 5.2.9. After gelation, the initial
weight of each hydrogel was determined. To study the degradation of the gel networks,
the samples were immersed in 10 mL of PBS (containing 0.025 % sodium azide) and
incubated at 37 ◦C in a shaking water bath (approx. 50 rpm). Every day, the samples
were poured out over 24 mm NetwellTM
Inserts (Corning GmbH, Kaiserslautern,
Germany) with a 500 µm mesh size polyester membrane and weighed. Degradation
was assumed to be complete when the sample passed through the NetwellTM
Insert
and no remaining material could be detected.
88
5.2 Materials and methods
5.2.11 Mobility of incorporated macromolecules determined byfluorescence recovery after photobleaching (FRAP)
To investigate the mobility of incorporated macromolecules with and without amino
groups, the hydrogels were loaded with FITC-BSA (66 kDa molecular weight) and
FITC-dextran (150 kDa molecular weight). The samples were prepared by dissolv-
ing 12.5 mg of 4armPEG10k-NH2 in 450 µL of 25 mM phosphate buffer, pH 7.4.
Subsequently, 50 µL of the FITC-BSA or FITC-dextran stock solution (prepared
in the same buffer, c = 10 mg/mL) were added. The mixture was then added to
12.5 mg of 4armPEG10k-dTyr-SC (degradable gels) or 12.5 mg of 4armPEG10k-SC
(non-degradable control gels), vigorously stirred, and cast into Lab-TekTM
II Cham-
bered Coverglasses (Thermo Fisher Scientific, Langenselbold, Germany). After 2 h of
gelation, the samples were positioned on the microscope stage.
The FRAP experiments were performed on a Zeiss Axiovert 200 M microscope
coupled to a LSM 510 META scanning device (Carl Zeiss MicroImaging GmbH, Jena,
Germany). A Plan-Neofluar 20× objective lens with a numerical aperture of 0.50 was
used. All bleaching experiments were performed using the 488 nm-line of a 30 mW
Ar-ion laser operating at 50 % output power. After the region of interest was brought
into focus, a time series of digital images with a resolution of 512 × 128 pixel was
recorded using a highly attenuated laser beam (0.5 % transmission). The interval
between two consecutive images was 2 s. After acquisition of five prebleach images,
a uniform disk with a diameter of 36 µm was bleached at maximum laser intensity
(100 % transmission). Immediately after bleaching, the laser intensity was switched
back to the previous value (0.5 % transmission), and a series of 75 images was acquired
in order to measure the recovery of fluorescence. To extract the experimental recovery
curve from the image stack, the mean fluorescence intensities inside the bleached
region, Ifrap(t), and inside a reference region were calculated for each time point using
the NIH software ImageJ. In the next step, Ifrap(t) was normalized to the prebleach
intensity, corrected for any bleaching effects that might have occurred during image
acquisition, and normalized to the full scale.
In order to follow changes in the macromolecular mobility during gel degradation,
the samples were covered with 500 µL of PBS (containing 0.025 % sodium azide) and
incubated at 37 ◦C. At predetermined time points (t = 48 h, 96 h, and 168 h), the
FRAP experiments were repeated as described above.
89
Chapter 5 Biodegradable hydrogels for time-controlled release
5.2.12 Release of FITC-BSA and lysozyme
The hydrogels were loaded with FITC-BSA (66 kDa molecular weight) and lysozyme
(14 kDa molecular weight) in order to evaluate their suitability as potential drug
delivery systems. These two proteins were chosen because of their different mass and
different number of amino groups, and serve as model compounds for therapeutic
peptides or proteins. To prepare the gel samples, 25 mg of 4armPEG10k-NH2 were
dissolved in 900 µL of 25 mM phosphate buffer, pH 7.4 and mixed with 100 µL of the
respective protein stock solution (prepared in the same buffer, c = 10 mg/mL). This
mixture was then added to 25 mg of 4armPEG10k-dTyr-SC and vigorously vortexed.
In each case, 250 µL of these precursor solutions were cast into cylindrical glass molds
(7 mm inner diameter) and allowed to gel for 2 h. The protein loading was 250 µg
per gel. Afterwards, the gels were removed from the glass molds, immersed in 10 mL
of PBS (containing 0.025 % sodium azide), and maintained at 37 ◦C in a shaking
water bath (approx. 50 rpm). All vials were treated with Sigmacote R© to prevent the
adsorption of proteins. At specified time intervals, 500 µL of the release medium were
collected and replaced with fresh buffer. Blank hydrogels without protein and protein
solutions (250 µg in 10 mL of PBS) served as control groups.
The released protein amounts were determined as described by Bradford [163].
In brief, 100 µL of sampled release buffer were pipetted into a microtiter plate and
incubated with 100 µL of Bradford reagent for 10 min. The protein content was
quantified by measuring the absorption at 595 nm using a Shimadzu CS-9301PC
96-well plate reader (Shimadzu GmbH, Duisburg, Germany). Calibration curves were
obtained from known concentrations of FITC-BSA and lysozyme. All experiments
were carried out in triplicate and the results are expressed as means ± standard
deviations.
5.2.13 Statistical analysis
The results from mechanical testing, swelling studies, and FRAP experiments were
compared using single-factor analysis of variance (ANOVA) and Tukey’s multiple
comparison test. p < 0.05 was regarded as statistically significant. Statistical analysis
was performed using SigmaStat 3.0 software (Systat Software, San Jose, CA).
90
5.3 Results and discussion
5.3 Results and discussion
The aim of the present study was to prepare in situ forming hydrogels that allow
for the time-controlled release of incorporated proteins. For this purpose, branched
PEG-succinimidyl carbonates were synthesized (Figure 5.2A). These polymers react
with primary amino groups of other polymers, peptides, or proteins under formation
of carbamate bonds. In case of proteins, succinimidyl carbonates typically react
with N-terminal amino acids and ε-amino groups of lysine residues [164]. Since bond
formation and cleavage may vary depending on the chemical environment of the
reacting amino groups, the polymers used for gel formation were further modified
with alanine (mimicking the amino terminus of proteins), 6-aminohexanoic acid, and
lysine (Figure 5.2B). All syntheses were straightforward and the desired products
were obtained with high yields. End-groups were almost fully converted, as indicated
by 1H-NMR spectra.
5.3.1 Physicochemical characterization of hydrogels
Hydrogels were prepared by step-growth polymerization of branched, amino acid-
modified PEG-amines with branched PEG-succinimidyl carbonates. As it was ex-
pected, gelation kinetics and gel strength were strongly dependent on the polymers
used for gel preparation. In preliminary tests, the buffer pH value was therefore
adjusted for each gel type (Table 5.1). When the buffer pH value was too low,
polymerization was slow and the gel strength remained comparatively low. On the
other hand, if the pH value was too high, the gels solidified immediately and could
no longer be placed on the rheometer.
Non-degradable hydrogels were prepared from 4armPEG10k-SC and 4armPEG10k-
NH2 at pH 7.0. At the beginning of the experiment, the sample behaved like a free
flowing liquid (G′′ > G′); gelation occurred after 9.2± 0.4 min. During the course of
the experiment, cross-linking further proceeded as indicated by the steadily increasing
value of G′ (Figure 5.3). After 90 min, the value of G′ exceeded that of G′′ by
several orders of magnitude (G′ � G′′); the gel strength was 2429± 73 Pa (Table 5.2).
These data agreed very well with those reported previously for gels prepared from
branched PEG-succinimidyl propionates and PEG-amines [139]. For the preparation
91
Chapter 5 Biodegradable hydrogels for time-controlled release
0 10 20 30 40 50 60
0.01
0.1
1
10
100
1000
10000
0.01
0.1
1
10
100
1000
10000
nG
'(P
a)l
Time (min)
Control (G' )
Control (G" )
PEG-NH2(G' )
PEG-NH2(G" )
¨G
"(P
a)¡
Figure 5.3: Typical rheograms of degradable (p/@) and non-degradable hydrogels(u/E). Both gels were prepared from 4armPEG10k-NH2 at pH 7.4 (degradable gel) andpH 7.0 (non-degradable gel). The closed symbols (p/u) represent G′, the open symbols(@/E) represent G′′.
of biodegradable gels, 4armPEG10k-NH2 was polymerized with 4armPEG10k-dTyr-
SC at pH 7.4. These samples solidified in less than 1 min and the gel strength
was 1486 ± 132 Pa (Table 5.2). Compared to 4armPEG10k-SC, the reactivity of
4armPEG10k-dTyr-SC seemed to be considerably increased (Figure 5.3). This could
explain both the accelerated polymerization as well as the lower strength of these
gels. With increasing reactivity of succinimidyl carbonate groups, their susceptibility
to hydrolysis will simultaneously increase [160]. This will result in lower cross-linking
densities and hence reduced gel strengths. In case of 4armPEG10k-dLys, the buffer
pH value was increased to pH 8.0. This was required because of the high pKa value of
the 6-aminohexanoic acid moiety (10.6± 0.1). The prepared hydrogels solidified after
1.1±0.2 min and the absolute value of G∗ was 1075±68 pascal (Table 5.2). However,
when 4armPEG10k-Ala and 4armPEG10k-Lys were used for gel preparation, the
buffer pH value was decreased to pH 6.8 and pH 6.4, respectively. Otherwise, gelation
occurred within a few seconds and the samples could not be placed on the rheometer.
The high reactivity of 4armPEG10k-Ala was attributed to the comparatively low pKa
value of the alanine moiety (8.2± 0.3). And in case of 4armPEG10k-Lys, gelation
92
5.3 Results and discussion
was most likely facilitated by the increased number of amino groups per macromer (8
vs. 4). The stiffness was 2137± 55 Pa for gels prepared from 4armPEG10k-Ala, and
2435± 105 Pa for those prepared from 4armPEG10k-Lys (Table 5.2).
Table 5.2: Gel strength (|G∗|), volumetric swelling ratio (Q), molecular weight betweencross-links (Mc), and average network mesh size (ξ) of the prepared hydrogels. Data arerepresented as means ± standard deviations (n = 3). The differences in the mean valuesare statistically significant (p < 0.05).
Group |G∗| (Pa) Q Mc (g ·mol−1) ξ (nm)
PEG-dLys 1075 ± 68 36.8 ± 0.5 10477 ± 202 21.2 ± 0.3
PEG-NH2 1486 ± 132 34.7 ± 0.1 9300 ± 82 19.6 ± 0.1
PEG-Ala 2137 ± 55 32.2 ± 0.3 7905 ± 123 17.6 ± 0.2
PEG-Lys 2435 ± 105a 30.9 ± 0.3b – –
Control 2429 ± 73a 30.6 ± 0.8b 6946 ± 122 16.3 ± 0.2
aStatistically not significant (p > 0.05)bStatistically not significant (p > 0.05)
When compared to each other, the gels prepared from 4armPEG10k-dLys had the
lowest cross-linking density of all degradable hydrogels (estimated by G∗), whereas
those made from 4armPEG10k-Lys showed the highest value. This corresponded
well with the results of equilibrium swelling studies. The volumetric swelling ratio
Q was inversely related to gel strength and was highest for hydrogels prepared from
4armPEG10k-dLys and lowest for those prepared from 4armPEG10k-Lys (Table 5.2).
Using Equations (5.1) and (5.2), these data also allowed for the calculation of the
average network mesh size (ξ). The gels with the lowest cross-linking density showed
the highest value of ξ (21.2±0.3 nm); hydrogels with higher stiffness and hence higher
cross-linking density exhibited lower values of ξ (Table 5.2). For gels prepared from
4armPEG10k-Lys, the average network mesh size could not be calculated, as these
gels contained macromers of two different functionalities and Equation (5.1) can only
be applied for gels containing one single type of cross-links [140–142].
93
Chapter 5 Biodegradable hydrogels for time-controlled release
5.3.2 Degradation of hydrogels
Samples were incubated in PBS at 37 ◦C and weighed at predetermined time points to
study hydrogel degradation. As expected, the gels of the control group did not degrade
within the observation period. During the first 24 h, the wet weight of these samples
first increased due to swelling. However, after the non-degradable gels were swollen
to equilibrium, their wet weight remained constant (Figure 5.4). In these networks,
the individual macromers are linked together by aliphatic carbamate bonds, which
are stable under the experimental conditions [165]. In degradable gels, however, the
polymer chains were held together by aromatic carbamate groups. These carbamates
hydrolyze in neutral and basic solutions by an E1cB elimination reaction involving the
intermediate formation of an unstable isocyanate [165, 166]. The isocyanate readily
reacts with water and disintegrates into a primary amine and carbon dioxide (see
Figure 5.1 for comparison) [165, 166].
0 12 24 36 48 60 72 84 96 108 120
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
Wetw
eig
ht(g
)
Time (h)
PEG-dLys
PEG-NH2
PEG-Ala
PEG-Lys
Control
Figure 5.4: Degradation of hydrogels. Gels prepared from 4armPEG10k-Lys (u) and4armPEG10k-Ala (p) degraded within 48 h. Hydrogels made from 4armPEG10k-NH2
(q) and 4armPEG10k-dLys (f) disintegrated over 4 and 5 days, respectively. The non-degradable gels (E) remained stable over the observation period. The experiments werecarried out in triplicate and the results are presented as means ± standard deviations.
94
5.3 Results and discussion
During the initial phase of the study, the wet weight of the degradable samples
first increased (Figure 5.4). The hydrolysis of carbamate groups obviously enlarged
the average network mesh size of these hydrogels which resulted in an increased
swelling capacity. After a critical amount of bonds had been cleaved, the wet weight
decreased and the gels dissolved slowly. Interestingly, the two gels prepared from
4armPEG10k-Lys and 4armPEG10k-Ala disintegrated within the first 48 h. This
was not expected, as these hydrogels had the highest cross-linking density of all
biodegradable gels. In contrast to that, the hydrogels containing 4armPEG10k-NH2
and 4armPEG10k-dLys were stable over 72 h and 96 h, respectively, and dissolved
within the following 24 h (Figure 5.4).
Obviously, the nature of the amino groups not only influenced gelation kinetics
and gel strength but also affected degradation rate. It is known from the literature
that the reactivity of phenyl carbamates increases with increasing polar effects within
the N-substituent. Phenyl carbamates of amino acid amides or dipeptides, for
example, were hydrolyzed within a few hours. Phenyl N-ethylcarbamates, however,
showed a half-life of several days [166]. The same effects could account for the
degradation profiles of the different hydrogels. In case of 4armPEG10k-Lys and
4armPEG10k-Ala, the amino group is in close proximity to an amide linkage, which
makes the resulting carbamates more susceptible to hydrolysis. In contrast to that,
the chemical structures of 4armPEG10k-NH2 and 4armPEG10k-dLys resemble those
of phenyl N-ethylcarbamates, which can explain the better hydrolytic resistance of
the corresponding hydrogels.
As a compromise between both, gel strength and degradation time, hydrogels
prepared from 4armPEG10k-NH2 were chosen for all further experiments. In future
studies it may be also possible to prolong the degradation time by introducing strongly
electropositive carboxylic acid moieties into the N-substituent, since these groups
were reported to increase the hydrolytic resistance of phenyl carbamates [166].
5.3.3 Mobility of incorporated macromolecules
To show their suitability as drug delivery system, the developed hydrogels were
loaded with FITC-BSA (66 kDa molecular weight) and FITC-dextrans (150 kDa
95
Chapter 5 Biodegradable hydrogels for time-controlled release
molecular weight). The hydrodynamic diameters of these two macromolecules were
7.2 nm [167] and 16.6 nm [168], respectively. When comparing these values with
the average network mesh sizes of non-degradable (16.3 nm) and degradable gels
(prepared from 4armPEG10k-NH2, 19.6 nm), the diffusivity of both macromolecules
should be restricted only to a minor extent. The diffusivity of the incorporated
FITC-dextran served as benchmark and allowed for the evaluation of the successful
immobilization of FITC-BSA.
A) B)
*
*
*
*
FITC-BSA FITC-Dextran
0 %
20 %
40 %
60 %
80 %
100 %
120 %
Mo
bile
fracti
on
Non-degradableDegradable
0 48 96 144 192
0 %
20 %
40 %
60 %
80 %
100 %
120 %
Time (h)
Mo
bile
fracti
on
Non-degradableDegradable
C)
0 20 40 60
0.0
0.2
0.4
0.6
0.8
1.0
1.2 t = 0 h t = 48 ht = 96 h t = 168 h
No
rmalize
dfl
uo
rescen
ce
inte
nsit
y
Time (s)
Figure 5.5: Mobile fractions of FITC-dextran and FITC-BSA in non-degradable (@)and degradable (p) hydrogels directly after cross-linking (A). * indicates statisticalsignificance (p < 0.05). Mobile fractions of FITC-BSA in non-degradable (E) anddegradable (u) hydrogels over time (B). Mobility of FITC-BSA in degradable hydrogels(C). The recovery profiles were acquired 0 h (u), 48 h (p), 96 h (q), and 168 h (f)after cross-linking.
Directly after cross-linking, 73 % of the FITC-dextran molecules incorporated
into non-degradable hydrogels were mobile. In case of degradable gels, even 98 %
of the incorporated FITC-dextrans were mobile (Figure 5.5A). Due to the lack of
96
5.3 Results and discussion
amino groups, FITC-dextrans cannot be bound to the gel network by reaction with
PEG-succinimidyl carbonates. The obtained results further indicate that incorporated
macromolecules will not be immobilized within the gel network by physical entrapment.
In FITC-BSA loaded hydrogels, however, the situation was different. Directly after
hydrogel preparation, most of the incorporated protein molecules were immobilized
within the gel network. The mobile fractions were 4 % in non-degradable gels, and
20 % in degradable hydrogels (Figure 5.5A). Since the hydrodynamic diameter of
FITC-BSA was well below the average network mesh size of the prepared gels, it
was concluded that the protein molecules were successfully immobilized by covalent
attachment to the hydrogel backbone. This demonstrates the general potential for
tethering therapeutic peptides or proteins to the gel backbone by simply dissolving
them together with the gel-forming polymers. Hydrogel cross-linking and drug loading
can be performed simultaneously without the need for additional synthetic steps.
To follow the mobility of FITC-BSA over time, the gel samples were covered with
PBS. In non-degradable hydrogels, protein mobility did not change substantially
during the observation period. After 168 h of incubation, approx. 20 % of the
incorporated protein molecules seemed to be mobile (Figure 5.5B). This increase can
be attributed to hydrolysis of aliphatic carbamate groups, slow degradation of the
protein, or release of the fluorescence label. In degradable hydrogels, however, FITC-
BSA mobility gradually increased (Figure 5.5B and C). After 168 h, when the hydrogel
was completely degraded, all protein molecules were mobile. The lower degradation
rate most likely results from differences in the experimental setup (see Figure 5.4 for
comparison). The FRAP experiments indicate that the developed hydrogels would
allow for the time-controlled release of therapeutic peptides or proteins. As a result
of the degradation mechanism, encapsulated peptides or proteins were expected to
be released in the unaltered state, which preserves bioactivity and lowers the risk of
unwanted immune reactions [169, 170].
5.3.4 Release of FITC-BSA and lysozyme
In the last experiment, the release of FITC-BSA and lysozyme was quantified. As
expected from degradation studies and FRAP experiments, almost no FITC-BSA was
released during the first 24 h (Figure 5.6). With the onset of gel degradation, however,
97
Chapter 5 Biodegradable hydrogels for time-controlled release
more and more protein was released into solution. The obtained release profile had a
sigmoidal shape and matched the degradation profile very well. After 96 h, the release
of FITC-BSA was completed. In FRAP experiments, only 75 % of the incorporated
protein molecules were mobile after the same time period. These differences are most
likely due to the different amounts of PBS used for FRAP and release experiments
(500 µL vs. 10 mL of PBS). Compared to FITC-dextrans, which were released from
similar gels almost completely within 24 h [139], the covalent attachment considerably
prolonged the release of incorporated protein molecules. In addition to FITC-BSA,
the hydrogels were also loaded with lysozyme. In general, the resulting release profile
was similar to that of FITC-BSA (Figure 5.6). During the initial phase, however, the
release of lysozyme was significantly higher. After 54 h, approx. 30 % of the total
amount of lysozyme was released into solution. In the case of FITC-BSA, however,
only 15 % of the incorporated protein molecules were released at the same time point.
Furthermore, the release profile of lysozyme was almost linear during the first 54 h.
0 24 48 72 96 120 144 168 192
0 %
20 %
40 %
60 %
80 %
100 %
120 %
Mt/M
0
Time (h)
FITC-BSALysozyme
Figure 5.6: Release of FITC-BSA (u) and lysozyme (E) from biodegradable hydrogels.Data are presented as means ± standard deviations (n = 3).
These variations are explained by the different characteristics of the encapsulated
proteins. In the case of lysozyme, one protein molecule can be bound to the gel
network by a maximum of 7 amino groups (amino terminus and ε-amino groups
98
5.4 Conclusion
of lysine residues). Bovine serum albumin, in contrast, is bearing 60 amino group
(amino terminus and ε-amino groups of lysine residues) that can theoretically react
with the hydrogel backbone. The probability that one protein molecule is detached
from the polymer network is therefore much higher for lysozyme than for FITC-BSA.
Furthermore, the lower mass of lysozyme (14 kDa vs. 66 kDa) results in an increased
diffusivity within the hydrogel network. This effect will be particularly pronounced
during the initial phase of protein release since the average network mesh size increases
during hydrogel degradation.
Altogether, the prepared hydrogels proved to be suitable for the time-controlled
release of incorporated peptides or proteins. The obtained release profiles will depend
on both the encapsulated macromolecules and the polymers used for gel formation.
In future experiments, different polymers (e.g. fast and slow degrading macromers)
could be combined to adjust the resultant release profiles. In the end, this might
allow for a constant release of therapeutic peptides or proteins over a time period of
several days.
5.4 Conclusion
We successfully synthesized different derivatives of poly(ethylene glycol) that allow for
the preparation of in situ forming hydrogels. Gel strength and degradability could be
tailored by altering the polymer end-groups. Since cross-linking is performed in situ,
the developed hydrogels could be easily delivered by injection. During the gelation
process, dissolved proteins were covalently bound to the polymer backbone, as shown
by FRAP experiments. In the same way, therapeutic peptides or proteins could be
tethered to the hydrogel network without the need for chemical modifications of these
molecules. The non-radical cross-linking approach is, thereby, favorable to the stability
of these fragile molecules. During hydrogel degradation, the incorporated proteins
were released into solution. Release kinetics will depend on both the incorporated
proteins and the polymers used for gel formation. It is expected that the chosen linker
group disintegrates without leaving any residues attached to the protein; this would
preserve bioactivity and lower the risk of unwanted immune reactions. Altogether,
99
Chapter 5 Biodegradable hydrogels for time-controlled release
the developed hydrogels proved to be suitable for the time-controlled release of
incorporated molecules. Further modifications of the described polymers might result
in long-lasting hydrogels that would allow for the sustained release of therapeutic
peptides or proteins over a time period of several days up to a few weeks.
100
Chapter 6
Biointeractive hydrogels for adiposetissue engineering
Ferdinand Brandl1, Annina Seitz1, Jorg Teßmar1, Torsten Blunk1,
Achim Gopferich1
1 Department of Pharmaceutical Technology, University of Regensburg, 93040 Regensburg
Submitted to Biomaterials.
101
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
Abstract
Adipose tissue engineering requires biomaterials that promote the differentiation of
seeded adipocytes. Here, we report on the development and characterization of in situ
forming, poly(ethylene glycol) (PEG) based hydrogels for soft tissue augmentation.
Branched PEG-amines were modified with collagenase-sensitive peptides and cross-
linked with branched PEG-succinimidyl propionates without the use of free-radical
initiators (enzymatically degradable hydrogels). Alanine-modified PEG-amines were
used for the preparation of non-degradable gels. Depending on the used polymer
concentration, the strength of degradable gels ranged from 1708 to 7412 Pa; the
strength of non-degradable hydrogels varied between 1496 and 7686 Pa. Enzyme
mediated gel degradation occurred within 10, 16, and 19 days (5 %, 10 %, and
15 % initial polymer content). To evaluate their suitability as scaffold materials for
adipose tissue engineering, the hydrogels were functionalized with the laminin-derived
adhesion peptide YIGSR, and seeded with 3T3-L1 preadipocytes. Compared to a
standard two-dimensional cell culture model, the developed hydrogels significantly
enhanced the intracellular triglyceride accumulation of encapsulated adipocytes.
Functionalization with YIGSR further enhanced lipid synthesis within differentiating
adipocytes. Long-term studies suggested that enzymatically degradable hydrogels
furthermore promote the formation of coherent adipose tissue-like structures featuring
many mature unilocular fat cells.
102
6.1 Introduction
6.1 Introduction
In reconstructive surgery, there is a tremendous need for suitable grafts to repair
soft tissue defects following tumor resections, for example. However, none of the
currently used strategies has proven to be ideal for permanent soft tissue augmentation.
Adipose tissue engineering has been proposed as an alternative approach to generate
functional tissue substitutes [171–174]. This will require biomaterials with mechanical
properties close to those of natural adipose tissue; very rigid scaffolds will certainly
not be appropriate to augment soft tissue defects. With regard to the desired
substrate mechanics, poly(ethylene glycol) (PEG) based hydrogels would be qualified
as three-dimensional (3-D) scaffolds for adipose tissue engineering [171, 172, 175–178].
These hydrophilic polymer networks absorb large amounts of water, allow for the
unrestricted diffusion of low molecular weight nutrients and metabolites [179], and
show an excellent biocompatibility because of their physicochemical similarity to
the native extracellular matrix (ECM) [10]. In addition to the initial mechanical
properties, the degradability of the supportive matrix must be considered as well.
Current tissue engineering strategies favor biodegradable scaffolds that provide only
transient stability; the newly developed tissue and the formed ECM will be responsible
for the long-term maintenance of the tissue engineered construct. Therefore, it will be
necessary to adapt the degradation rate to the rate of tissue formation [8, 10]. Since
most synthetic hydrogels are biologically inert due to their limited interactions with
ECM proteins, the gel-forming polymers may additionally be functionalized with cell
adhesion ligands or growth factors in order to promote cell migration, proliferation,
and differentiation [8, 10, 15].
For the preparation of such ‘biomimetic’ hydrogels, Lutolf et al. cross-linked vinylsul-
fone terminated PEG macromers with cysteine containing, matrix metalloproteinase
(MMP) sensitive peptides [74, 90, 180]. In contrast to biomaterials with hydrolyti-
cally labile bonds, such as poly(lactic acid) blocks for example, these cell-responsive
hydrogels are degraded by cell-secreted and cell-activated proteases. The gels formed
in situ, allowed for the invasion of cells, and proved to be suitable for the regeneration
of bone [100] and cartilage defects [82]. In another approach, collagenase-sensitive
peptides (Gly–Gly–Leu↓Gly–Pro–Ala–Gly–Gly–Lys) and integrin-binding domains
(Tyr–Ile–Gly–Ser–Arg) were coupled with amine-reactive PEG-monoacrylates. The
103
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
resulting triblock copolymers were terminated by acrylate groups and allowed for the
preparation of photopolymerizable hydrogels that supported viability, adhesion, and
proliferation of preadipocytes [176].
But despite these promising results, non-radical cross-linking schemes will be
favored for the encapsulation of cells [181]. Furthermore, cytotoxic byproducts
(such as unreacted macromers or degradation products of the polymers) may be
leached out of radically cross-linked hydrogels, which have to be removed prior
to their implantation into the patient [182]. Therefore, we aimed for developing
injectable, biointeractive hydrogels that are cross-linked without the use of free-radical
initiators (Figure 6.1). For this purpose, branched PEG-amines were functionalized
with the collagenase-sensitive amino acid sequence Ala–Pro–Gly↓Leu (cleavage site
between glycine and leucine) by means of classical liquid phase peptide synthesis.
For the formation of hydrogels, these polymers were cross-linked with branched,
amine-reactive PEG-succinimidyl propionates. In contrast to previously reported
approaches, the described hydrogels can be functionalized with adhesion peptides or
growth factors without requiring additional chemical modifications of these molecules
(e.g. introduction of acrylate groups or cysteine residues). The prepared hydrogels were
characterized for their mechanical properties, swelling behavior, and degradability. To
evaluate their potential as scaffold materials for soft tissue engineering, these gels were
seeded with 3T3-L1 preadipocytes, a commercially available cell line that has been
extensively used to study adipocyte differentiation in vitro [174, 183]. By quantifying
the intracellular triglyceride accumulation, we studied the effects of substrate stiffness
and adhesiveness on adipocyte differentiation, and compared these results to the
outcome of conventional two-dimensional (2-D) cell culture. Finally, we showed the
advantage of enzymatically degradable hydrogels for the in vitro generation of adipose
tissue-like constructs.
6.2 Materials and methods
6.2.1 Materials
Hexane, methylene chloride (DCM), and phthalimide was purchased from Acros
Organics (Geel, Belgium). Murine 3T3-L1 preadipocytes were obtained from ATCC
104
6.2 Materials and methods
NH
O
NH
NH
O
O
N
O
NH
O
Adhesion peptideHO-Arg-Ser-Gly-Ile-Tyr–
Collagenase substrate
–Leu Gly-Pro-Ala–¯
C
B
A
NH
NH
NH
NH
NH
O
O
O
O
O
OH
O
OH
OH
NH
NH
NH2
O O
O
O N
O
O
OO N
H
N
O
OO
NH
+
N
O
O
OH
NH2
N
OO
NH
Figure 6.1: Schematic illustration of injectable, biointeractive hydrogels for soft tissueengineering applications. For gel formation, branched PEG-succinimidyl propionatesare cross-linked with branched, amino acid-modified PEG-amines (A). The collagenase-sensitive peptide sequence Ala–Pro–Gly↓Leu allows for enzymatic gel degradation (B).To mediate cell-adhesion, the integrin-binding motif Tyr–Ile–Gly–Ser–Arg is reacted withbranched PEG-succinimidyl propionates (C).
(Manassas, VA). Boc-protected alanine (Boc-Ala-OH), Boc-protected glycine (Boc-
Gly-OH), and Boc-protected proline (Boc-Pro-OH) were bought from Bachem GmbH
(Weil am Rhein, Germany). Dulbecco’s modified Eagle’s medium (DMEM) and
trypsin (1 : 250) were purchased from Biochrom (Berlin, Germany). Deuterated
chloroform (CDCl3) was obtained from Deutero GmbH (Kastellaun, Germany). Fetal
bovine serum (FBS, Lot. No. 40A0044K), penicillin-streptomycin solution, and
phosphate buffered saline (PBS) were from Invitrogen GmbH (Karlsruhe, Germany).
Four-armed poly(ethylene glycol) (molecular weight 10 kDa, 4armPEG10k-OH) was
purchased from Nektar Therapeutics (Huntsville, AL). Hoechst 33258 dye was obtained
from Polysciences (Warrington, PA); calcium chloride (CaCl2) was from Carl Roth
GmbH (Karlsruhe, Germany). Bovine insulin was kindly provided by Sanofi-Aventis
(Frankfurt am Main, Germany). 3-Isobutyl-methylxanthine (IBMX) was bought
from Serva Electrophoresis (Heidelberg, Germany). Acrylonitrile, Boc-protected
leucine (Boc-Leu-OH), collagenase from Clostridium histolyticum (Type I), corticos-
105
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
terone, cysteine, N,N’ -dicyclohexylcarbodiimide (DCC), diisopropyl azodicarboxy-
late, N,N’ -diisopropylethylamine (DIPEA), DNA from calf thymus, fluorescamine,
N-hydroxysuccinimide (HOSu), indomethacin, the laminin fragment Cys–Asp–Pro–
Gly–Tyr–Ile–Gly–Ser–Arg (YIGSR), Minimum Essential Medium alpha modification
(α-MEM), oil red O, and Sigmacote R© were purchased from Sigma-Aldrich (Taufkirchen,
Germany). Papainase was bought from Worthington (Lakewood, NJ). Diethyl ether
was of technical grade and used without further purification. All other chemicals
were of analytical grade and obtained from Merck KGaA (Darmstadt, Germany).
Deionized water was obtained using a Milli-Q water purification system from Millipore
(Schwalbach, Germany).
6.2.2 Synthesis of amine-reactive polymers (4armPEG10k-SPA)
Branched PEG-succinimidyl propionates (4armPEG10k-SPA) were synthesized as
previously described [139]. In brief, PEG-propionic acid was obtained through
Michael-type addition reaction of 4armPEG10k-OH onto acrylonitrile and subsequent
hydrolysis under alkaline conditions. The obtained carboxylic acid groups were then
converted into amine-reactive succinimidyl ester groups using HOSu and DCC.
6.2.3 Synthesis of collagenase-sensitive polymers(4armPEG10k-LGPA)
The synthesis of collagenase-sensitive polymers started from branched PEG-amines
(4armPEG10k-NH2), which were obtained as previously described [139]. In the
first step, phthalimido-PEGs were synthesized by reaction of 4armPEG10k-OH,
phthalimide, triphenylphosphine, and diisopropyl azodicarboxylate. The phthalimido
groups were then converted into primary amines by hydrazinolysis. In the next
step, the collagenase-sensitive tetrapeptide Ala–Pro–Gly↓Leu was synthesized using
4armPEG10k-NH2 as polymeric support [184]. For the coupling reaction, 5.55 g of
Boc-Leu-OH (24.0 mmol) and 2.48 g of DCC (12.0 mmol) were dissolved in 50 mL of
dried DCM. After stirring for 30 min at room temperature, the solution was passed
through a glass filter funnel to remove precipitated N,N’ -dicyclohexylurea (DCU), and
106
6.2 Materials and methods
combined with a solution of 4armPEG10k-NH2 (15.0 g, 1.5 mmol) in DCM (150 mL).
Then, 2000 µL of DIPEA (12.0 mmol) were added, and the mixture was stirred for
1.5 h at room temperature. The solution was concentrated under reduced pressure and
placed into an ice bath. The intermediate (4armPEG10k-Leu-Boc) was crystallized
at 0 ◦C under vigorous stirring by drop-wise addition of cold diethyl ether. The
precipitate was collected by filtration, washed with cold diethyl ether, and dried under
vacuum to yield 16.0 g (98 %). Thin layer chromatography (3 : 1 : 1 glacial acetic
acid/1-butanol/water) indicated the absence of free Boc-Leu-OH.1H-NMR (4armPEG10k-Leu-Boc, CDCl3, 300 MHz): δ 0.94 ppm (d, 24H, –CH2CH-
(CH3)2), 1.40 – 1.50 ppm (m, 4H, –CH2CH(CH3)2), 1.44 ppm (s, 36H, –OtBu),
1.60 – 1.70 ppm (m, 8H, –CH2CH (CH3)2), 3.41 ppm (s, 8H, R3CCH2O–), 3.65 ppm
(s, –OCH2CH2O–), 4.11 ppm (4H, –CH (NHBoc)R’), 5.02 ppm (4H, –CH(NH Boc)R’),
6.63 ppm (4H, –NH C(O)–)
The successful coupling of the amino acid was confirmed using a modified fluo-
rescamine assay [185]. In brief, 5.0 µmol of 4armPEG10k-Leu-Boc were dissolved in
500 µL of 50 mM borate buffer, pH 8.5 (c = 10 µmol/mL). As reference, 5.0 µmol
of 4armPEG10k-NH2 were dissolved in 500 µL of the same buffer and diluted to a
final concentration of 0.1 µmol/mL. Then, 50 µL of each sample were diluted with
1350 µL of 50 mM borate buffer, pH 8.5, and mixed with 600 µL of a fluorescamine
solution (15 mg in 50 mL of acetone). After vigorous stirring, the fluorescence in-
tensities (λex = 390 nm, λem = 480 nm) were measured on a PerkinElmer LS 55
Fluorescence spectrometer (Perkin-Elmer, Wiesbaden, Germany). The degree of
end-group conversion was determined to be 99.7 %.
To deprotect the amino acid moiety, 16.0 g of 4armPEG10k-Leu-Boc (1.5 mmol)
were dissolved in 176 mL of methanolic HCl (prepared by dropping 16 mL of acetyl
chloride into 160 mL of ice-cooled methanol). After stirring for 30 min at room
temperature, the solvent was evaporated, and the residue was dissolved in 80 mL of
water. The pH of the aqueous solution was adjusted with sodium hydroxide (3 M)
to 9 – 10. The raw product was subsequently extracted with DCM. The combined
organic phases were dried over anhydrous Na2SO4, and the solution was concentrated
under reduced pressure. The product was crystallized at 0 ◦C under vigorous stirring
by drop-wise addition of diethyl ether. The precipitate was collected, washed with cold
diethyl ether, and dried under vacuum to yield 14.9 g (97 %) of 4armPEG10k-Leu.
107
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
1H-NMR (4armPEG10k-Leu, CDCl3, 300 MHz): δ 0.85 – 1.10 ppm (m, 24H,
–CH2CH(CH3)2), 1.55 – 1.75 ppm (m, 8H, –CH2CH(CH3)2), 1.75 – 1.95 ppm (m,
4H, –CH2CH (CH3)2), 3.41 ppm (s, 8H, R3CCH2O–), 3.65 ppm (s, –OCH2CH2O–),
8.45 ppm (t, 4H, –NH C(O)–)
The described steps were repeated for the subsequent addition of Boc-Gly-OH,
Boc-Pro-OH, and Boc-Ala-OH. At each step, 1H-NMR spectra were recorded, and
the successful coupling of the amino acids was confirmed as described above. The
degree of end-group conversion was almost quantitative. The resulting material is
denoted 4armPEG10k-LGPA. The C-termini of the peptide (leucine residues) are
conjugated to the PEG chain, and the N-termini (alanine residues) are free.1H-NMR (4armPEG10k-LGPA, CDCl3, 600 MHz): δ 0.85 – 1.00 ppm (m, 24H,
–CH2CH(CH3)2), 1.34 ppm (d, 12H, –CH3), 1.50 – 1.74 ppm (m, 12H, –CH2CH (CH3)2),
1.90 – 2.25 ppm (m, 16H, –C(O)CH(R’)CH2CH2–), 3.41 ppm (s, 8H, R3CCH2O–),
3.65 ppm (s, –OCH2CH2O–), 3.83 ppm (4H, –C(O)CH (R’)–), 4.28 ppm (4H,
–C(O)CH (R’)–), 4.42 ppm (4H, –C(O)CH (R’)–), 4.48 ppm (4H, –C(O)CH (R’)–)
6.2.4 Synthesis of non-degradable polymers (4armPEG10k-Ala)
4armPEG10k-Ala was synthesized from 4armPEG10k-NH2 and Boc-Ala-OH in 91 %
yield as described for 4armPEG10k-LGPA. Non-degradable and degradable polymers
are both terminated by alanine residues; consequently, their reactivity towards PEG-
succinimidyl propionates will be comparable.1H-NMR (4armPEG10k-Ala, CDCl3, 600 MHz): δ 1.34 ppm (d, 12H, –CH(NH2)-
CH3), 3.41 ppm (s, 8H, R3CCH2O–), 3.64 ppm (s, –OCH2CH2O–), 7.55 ppm (t, 4H,
–NH C(O)–)
6.2.5 Rheological characterization of hydrogels
Gelation kinetics and gel strength were studied by performing oscillatory shear
experiments on a TA Instruments AR 2000 rheometer (TA Instruments, Eschborn,
Germany) with parallel plate geometry at 37 ◦C. For the preparation of hydrogels,
accurately weighed amounts of 4armPEG10k-LGPA (degradable gels) or 4armPEG10k-
108
6.2 Materials and methods
Ala (non-degradable gels) were dissolved in 1000 µL of 25 mM phosphate buffer, pH 7.4,
and cooled to 5 ◦C. Immediately before starting the experiment, the polymer solutions
were added to accurately weighed amounts of 4armPEG10k-SPA. The stoichiometric
ratio between succinimidyl propionate and amino groups was balanced, and the
overall polymer concentrations were 5 %, 10 %, and 15 % (w/v). After vortexing,
the precursor solutions were poured onto the bottom plate of the rheometer. The
upper plate (20 mm in diameter) was lowered to a gap size of 1000 µm, and the
measurement was started. The evolution of storage (G′) and loss moduli (G′′) was
recorded as a function of time at 1 Hz oscillatory frequency and a constant strain of
0.05. A solvent trap was used to prevent the evaporation of water. The cross-over
of G′ and G′′ was regarded as the gel point. After 60 min, the absolute value of
the complex shear modulus (G∗) was determined as a measure for the obtained gel
strength. The gel disks were then removed from the rheometer, immersed in 10 mL of
PBS, and incubated at 37 ◦C for 24 h. After swelling, the samples were again placed
on the lower plate, and the measuring gap was slowly closed until a normal force of
approx. 750 mN was reached. The resulting compression of the gels was sufficient to
prevent slippage. The following frequency sweep was conducted at constant strain
(0.01) as a function of frequency (from 0.1 to 10 Hz). Finally, an amplitude sweep
(from 0.1 to 10, 000 Pa oscillatory stress) was performed in order to confirm that all
previous measurements were conducted within the linear viscoelastic region of the
sample. All experiments were carried out in triplicate and the presented results are
expressed as means ± standard deviation.
6.2.6 Equilibrium swelling of hydrogels
For the swelling studies, accurately weighed amounts of 4armPEG10k-LGPA (degrad-
able gels) or 4armPEG10k-Ala (non-degradable gels) were dissolved in 1000 µL of
25 mM phosphate buffer, pH 7.4, cooled to 5 ◦C, and added to accurately weighed
amounts of 4armPEG10k-SPA. The stoichiometric ratio between succinimidyl propi-
onate and amino groups was balanced, and the overall polymer concentrations were
5 %, 10 %, and 15 % (w/v). After vortexing, 250 µL of the precursor solutions were
cast into cylindrical glass molds (7 mm inner diameter) and allowed to gel for 2 h
109
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
at room temperature. The gel samples were then weighed in air and hexane after
swelling for 24 h in 10 mL of PBS using a density determination kit (Mettler-Toledo,
Gieen, Germany). Based on Archimedes’ principle, the gel volume after swelling (Vgs)
was determined. The volume of the dry polymer (Vp) was calculated from the mass
of the freeze-dried hydrogel and the density of the polymer in the dry state (taken as
the density of PEG, 1.12 g · cm−3). With these parameters, the volumentric swelling
ratio, Q = Vgs/Vp, was calculated.
6.2.7 Degradation of hydrogels
The gel samples were prepared as described in section 6.2.6. Directly after gelation,
the initial weight of the hydrogels was determined. The gel cylinders were then
placed into 5 mL of 10 mM HEPES buffer, pH 7.4, which also contained 1 mM CaCl2
and 0.025 % sodium azide. The samples were incubated for 24 h at 37 ◦C to allow
them to reach their equilibrium swelling levels. Afterwards, their wet weight was
determined (day 0). To study gel degradation, the samples were immersed in 5 mL
of HEPES buffer and incubated at 37 ◦C in a shaking water bath (approx. 50 rpm),
either without enzyme, or with 0.01 mg/mL collagenase (collagen digestion activity:
196 U/mg solid). At predetermined time points, the samples were poured out over
24 mm NetwellTM
Inserts (Corning GmbH, Kaiserslautern, Germany) with a 500 µm
mesh size polyester membrane. The wet weight of the gel cylinders was determined,
and the buffer or enzyme solution was replaced. Degradation was assumed to be
complete when the sample passed through the NetwellTM
Insert and no remaining
material could be detected.
6.2.8 Cell seeding and cell culture
3T3-L1 preadipocytes were plated in T150 culture flasks (Corning, Bodenheim,
Germany) and expanded in DMEM supplemented with 10 % fetal bovine serum
(FBS), penicillin (100 U/mL), and streptomycin (100 µg/mL). Cell cultures were
incubated in a humidified atmosphere at 37 ◦C and 5 % CO2. The culture medium
110
6.2 Materials and methods
was exchanged every two days. At confluency, the 3T3-L1 cells were harvested using
trypsin-EDTA, centrifuged to a pellet (1200 rpm, 5 min) and washed with PBS.
For the preparation of hydrogels, accurately weight amounts of 4armPEG10k-
LGPA (degradable gels) or 4armPEG10k-Ala (non-degradable gels) were dissolved in
phosphate buffer (50 mM, pH 7.4) at 125 mg/mL. A solution of Cys–Asp–Pro–Gly–
Tyr–Ile–Gly–Ser–Arg (YIGSR, c = 10 µmol/mL) or phosphate buffer was added, so
that the final polymer concentrations were 100 mg/mL. In a similar manner, accu-
rately weight amounts of 4armPEG10k-SPA were dissolved in water at 100 mg/mL.
All polymer solutions were kept on ice until use. For cell seeding, the prepared
3T3-L1 preadipocytes were suspended in the amino component (4armPEG10k-LGPA
or 4armPEG10k-Ala). Subsequently, an equal volume of 4armPEG10k-SPA was
added. After vortexing, 50 µL of the polymer-cell solution were cast into cylindrical
glass molds (5 mm inner diameter) that had been treated with Sigmacote R©. The
final polymer concentration was 10 % (w/v) and the cell number 100,000 cells per
hydrogel construct. For long-term cell culture 5,000,000 cells were used. After 30 min
of gelation at 37 ◦C, the glass rings were removed. Each gel sample was transferred
into one well of 24-well plates and covered with 2 mL of basal medium (α-MEM
supplemented with 10 % FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin).
For standard 2-D cell culture, 100,000 3T3-L1 cells were seeded in 24-well plates and
covered with 2 mL of basal medium.
Two days after cell seeding, adipogenesis was induced by applying induction medium
(basal medium supplemented with 1 µM insulin, 0.1 µM corticosterone, 0.5 mM IBMX,
and 60 µM indomethacin). The time point of induction was referred to as day 0. At
day 2, the induction medium was replaced by differentiation medium (basal medium
supplemented with 1 µM insulin). The hydrogel constructs were dynamically cultured
(approx. 60 rpm) on an orbital shaker (Heidolph Instruments, Schwabach, Germany) in
a humidified atmosphere at 37 ◦C and 5 % CO2. The culture medium was exchanged
every two days.
111
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
6.2.9 Quantitative analysis of intracellular triglycerideaccumulation
Nine days after induction, the intracellular triglyceride content was quantified using a
serum triglyceride determination kit (Sigma-Aldrich, Taufkirchen, Germany). After
washing with PBS, the hydrogel constructs were harvested in 0.5 % thesit solution,
disintegrated using a micropistill (Hartenstein Laborbedarf, Wurzburg, Germany),
and sonicated (Branson Ultrasonic Corporation, Danbury, CT). The spectroscopic
quantification of triglycerides was performed according to the manufacturer’s instruc-
tions. All measurements were done in three biological replicates. The amount of
triglycerides per hydrogel construct was calculated and normalized to the cell number
of the respective sample as determined by the DNA assay.
6.2.10 DNA assay
Aliquots of disintegrated and sonicated hydrogel constructs were completely digested
with papainase (3.2 U/mL in 0.1 M Na2HPO4 buffer, pH 6.5 containing 10 mM
Na2EDTA and 2.5 mM cysteine) for 16 h at 60 ◦C, and the DNA content was
determined using the intercalating Hoechst 33258 dye (0.1 µg/mL in 0.1 M NaCl
containing 1 mM Na2EDTA and 10 mM Tris, pH 7.4) [186]. Fluorescence intensities
were determined at 365 nm excitation wavelength and 458 nm emission wavelength
on a LS 55 Fluorescence spectrometer (PerkinElmer, Wiesbaden, Germany) and
correlated to DNA contents using standard dilutions of double-stranded DNA (from
calf thymus). Cell numbers were calculated with a conversion factor of 26.1 pg of
DNA per cell, which was determined previously.
6.2.11 Oil red O staining
In order to visualize cytoplasmic lipid droplets, the hydrogel constructs were washed
with PBS, fixed in formalin (10 % in PBS) overnight, and stained with oil red O
(3 mg/mL in 60 % isopropanol) for 12 h [187]. Excessive dye was removed by washing
three times with PBS. Microscopical brightfield pictures were acquired at 20× magni-
112
6.3 Results and discussion
fication using a Zeiss Axiovert 200 M microscope equipped with a Zeiss AxioCam HRc
camera and the software ZEN 2008 (Carl Zeiss MicroImaging GmbH, Jena, Germany).
6.2.12 Statistics
All quantitative results are presented as mean values ± standard deviation. The
results of the triglyceride and DNA assay were compared using the unpaired Student’s
t-test. p < 0.05 was regarded as statistically significant. Statistical analysis was
performed using PASW Statistics 17.0 software (SPSS Inc., Chicago, IL).
6.3 Results and discussion
6.3.1 Physicochemical characterization of hydrogels
For the preparation of biodegradable hydrogels, branched PEG-amines were function-
alized with the synthetic tetrapeptide Ala–Pro–Gly↓Leu (4armPEG10k-LGPA). For
non-degradable hydrogels, alanine-modified PEG-amines (4armPEG10k-Ala) were
used instead of 4armPEG10k-LGPA. Gels were formed by step-growth polymerization
of these macromers with branched PEG-succinimidyl propionates (4armPEG10k-SPA).
At the beginning of the experiments, all samples behaved like free-flowing liquids
(G′′ > G′). In this state, the gels could be cast into molds and would easily pass
through small-gauge needles in order to be injected into soft tissue defects. Further-
more, cells could be suspended in the liquid precursor solutions, which ensured their
homogeneous distribution throughout the gel network. Although PEG-succinimidyl
esters may also react with amino groups of cell surface proteins, these polymers were
shown to be non-toxic and rapidly excluded from the cell surface without being taken
up [188]. Gelation generally occurred within minutes with only minor differences
between the tested experimental groups (Table 6.1). During the course of the experi-
ment, cross-linking further proceeded as indicated by the steadily increasing value of
G′. After approx. 60 min, the value of G′ reached a plateau and exceeded that of G′′
by several orders of magnitude (G′ � G′′). The gel strength (|G∗|) of the biodegrad-
113
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
able hydrogels ranged between 2230 ± 108 Pa (5 % polymer) and 9625 ± 407 Pa
(15 % polymer, Table 6.1). Comparable values were obtained for the non-degradable
hydrogels. In this group, the absolute value of G∗ ranged between 1849 ± 26 Pa
(5 % polymer) and 10150 ± 87 Pa (15 % polymer, Table 6.1). These data agreed
well with those reported previously for gels prepared from 4armPEG10k-SPA and
4armPEG10k-NH2 [139].
After swelling in PBS, G′ and G′′ were measured at constant strain as a function of
frequency. The prepared hydrogels showed an elastic response (G′ > G′′) within the
studied frequency range. Furthermore, both moduli were almost insensitive to the
oscillatory frequency, which is typical for covalently cross-linked networks [149]. In the
swollen state, the gel strength (determined at a frequency of 1 Hz) was generally lower
than in the non-swollen state, which can be attributed to the uptake of a large amount
of water (Table 6.1). Again, comparable values were obtained for degradable and
non-degradable hydrogels. Amplitude sweep experiments indicated that all previous
measurements were run well within the linear viscoelastic region of the samples.
When compared to each other, the gels prepared from 5 % polymer had the lowest
cross-linking density (estimated by G∗), whereas those made from 15 % polymer
showed the highest value. This corresponded well with the results of equilibrium
swelling studies. The volumetric swelling ratio (Q) showed the opposite trend than
G∗ and was highest for hydrogels prepared from 5 % polymer and lowest for those
prepared from 15 % polymer (Table 6.1). For that reasons, network defects seem to
play a significant role in the structure of the prepared hydrogels [74]. At low precursor
concentrations (5 % polymer), the concentration of elastically active chains is probably
decreased by non-reacted groups and loops. At high precursor concentrations (above
10 % polymer), these effects are less pronounced and Q decreased only slightly when
increasing the initial polymer concentration from 10 % to 15 % (Table 6.1).
With regard to their physicochemical characteristics, the prepared hydrogels meet
the basic requirements for a successful application in adipose tissue engineering.
They will be capable of being gelled in the presence of cells, allow for the unre-
stricted diffusion of nutrients and metabolites, and provide sufficient mechanical
stability to withstand the occurring mechanical loads. The obtained gel strength,
thereby, mainly depends on the initial polymer concentration; the different polymers
(4armPEG10k-LGPA or 4armPEG10k-Ala) only have a minor influence on the result-
114
6.3 Results and discussion
Tab
le6.
1:M
echa
nica
lpro
pert
ies
and
swel
ling
beha
vior
ofth
epr
epar
edhy
drog
els.
Dat
aar
epr
esen
ted
asm
eans±
stan
dard
devi
atio
n(n
=3)
.
Deg
rad
able
(4ar
mP
EG
10k-L
GP
A)
Non
-deg
rad
able
(4ar
mP
EG
10k-A
la)
Pol
ymer
conc
entr
atio
n5
%10
%15
%5
%10
%15
%
Gel
atio
nti
me
(min
)1.
8±
0.2
1.6±
0.1
1.4±
0.1
2.9±
0.0
1.5±
0.2
1.2±
0.1
Gel
stre
ngth|G∗ |
afte
rcr
oss-
linki
ng(P
a)22
30±
108
6805±
123
9625±
407
1849±
2663
75±
487
1015
0±
87
Gel
stre
ngth|G∗ |
afte
rsw
ellin
g(P
a)17
08±
307
5567±
461
7412±
263
1496±
110
5097±
137
7686±
216
Vol
ume
incr
ease
(%)
73±
114
1±
119
4±
171±
013
2±
117
6±
2
Vol
umet
ric
swel
ling
rati
oQ
29.7±
0.2
20.9±
0.3
18.2±
0.2
31.1±
0.1
22.0±
0.2
18.5±
0.2
115
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
ing gel. Therefore, biodegradable and non-degradable gels will have almost the same
initial mechanical properties, which is relevant for the subsequent cell culture studies.
On the other hand, the specific characteristics of adipocytes must be considered as well.
These cells enlarge to over 100 µm in diameter during maturation and constantly
change their size because of their lipid storage function [172]. For these reasons,
hydrogels of intermediate stiffness (above 10 % polymer content) may be especially
promising. These gels are mechanically stable to withstand the manipulations dur-
ing cell culture and expected to enable the full differentiation of preadipocytes. In
addition, their mechanical properties resemble those of human adipose tissue [176].
6.3.2 Degradation of hydrogels
Mechanical strength and degradability of the supportive matrix have been recognized
to have a strong influence on cell migration, proliferation, and differentiation [102].
In most tissue engineering strategies, the scaffold materials are designed to provide
only transient stability; the developing tissue will be responsible for the long-term
maintenance of the construct. Therefore, it is necessary to adjust the degradation
kinetics to the rate of tissue formation [172]. For this purpose, the gel-forming
polymers were functionalized with the synthetic tetrapeptide Ala–Pro–Gly↓Leu to
make them susceptible to proteolytic breakdown. To study gel degradation in vitro,
the hydrogel samples were swollen to equilibrium and incubated in PBS containing
0.01 mg/mL collagenase. Although the enzymatic activity should be preserved in the
presence of Ca 2+-ions [189], the collagenase solution was replaced every day.
After the non-degradable hydrogels had reached their equilibrium swelling levels
(day 0), their wet weight remained almost constant (Figure 6.2A). This effect was
independent of the absence or presence of collagenase, which illustrates the requirement
of the Ala–Pro–Gly↓Leu sequence for gel degradation. Conversely, these experiments
also showed the necessity of collagenase for the degradation of hydrogels. Without
enzyme, the wet weight of enzymatically degradable gel samples did not change
significantly during the course of the experiment. However, in the presence of
0.01 mg/mL collagenase, their wet weight considerably increased during the first 4
to 5 days. In the following days, gel decomposition further proceeded; after approx.
116
6.3 Results and discussion
10 days, the degradation process was completed (Figure 6.2A). It can be concluded
from these experiments that hydrogel degradation was due to the proteolytic activity of
collagenase; non-enzymatic hydrolysis of amide bonds did not significantly contribute
to gel degradation.
A)
B)
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
0 1 2 3 4 5 6 7 8 9 10 11
Time (d)
We
tw
eig
ht
(g)
5 % Polymer, non-degradable, 0.01 mg/ml collagenase
5 % Polymer, non-degradable, without collagenase
5 % Polymer, degradable, 0.01 mg/ml collagenase
5 % Polymer, degradable, without collagenase
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
1.6
1.8
0 2 4 6 8 10 12 14 16 18 20
Time (d)
We
tw
eig
ht
(g)
5 % Polymer, degradable, 0.01 mg/ml collagenase
10 % Polymer, degradable, 0.01 mg/ml collagenase
15 % Polymer, degradable, 0.01 mg/ml collagenase
Figure 6.2: Wet weight of non-degradable (E/@) and degradable (u/p) hydrogels inthe presence (E/u) and absence (@/p) of collagenase (A). Degradation of hydrogelscontaining 5 % (u), 10 % (p), and 15 % (q) polymer in the presence of 0.01 mg/mLcollagenase (B).
117
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
Comparable results were obtained for all tested polymer concentrations (5 %, 10 %,
and 15 % initial polymer content). During the initial phase of degradation, the gel
samples considerably swelled as some of the Ala–Pro–Gly↓Leu sequences were cleaved
by the action of collagenase (Figure 6.2B). This, in turn, enlarged the average network
mesh size of hydrogels and increased their water absorption capacity. After a critical
amount of peptide sequences had been degraded, the ongoing polymer loss was no
longer overcompensated by swelling, and the wet weight reached a plateau. This effect
was particularly pronounced in gels prepared from 10 % and 15 % polymer. After
the maximum weight had been exceeded, the mass of the samples linearly decreased
by ongoing release of polymer chains or gel fragments (Figure 6.2B). This indicates
that both, surface and bulk erosion contribute to gel degradation. Gel degradation
was completed after 10, 16, and 19 days (5 %, 10 %, and 15 % initial solid content).
These findings correlated well with the results of equilibrium swelling studies. As
expected, the gels with the lowest cross-linking density (5 % initial polymer content,
Q = 29.7) showed the lowest resistance in the presence of collagenase (10 days
degradation time). In contrast to that, the gels prepared from 10 % and 15 % polymer
had comparatively high cross-linking densities (Q = 20.9 and 18.2, respectively).
Consequently, these hydrogels showed an increased stability towards proteolytic
degradation with only moderate differences between the two polymer concentrations
(16 and 19 days degradation time).
These results clearly demonstrate the enzyme dependent degradability of the
synthesized polymers. Transferring these data to the situation in cell culture will be
difficult, however. Here, the observed degradation rate will strongly depend on the
local concentration of proteolytic enzymes, which may vary over time [180, 190, 191].
The expression pattern of cellular proteases will also depend on whether the cells
are proliferating or differentiating, and their ability to migrate within the hydrogel
network [192]. Although the exact degradation rate can hardly be predicted, this
degradation mechanism is still favored over spontaneous hydrolytic degradation
since material resorption is caused and affected by cellular activity. As a result,
the degradation rate of the scaffold will be more closely related to the rate of
tissue formation.
118
6.3 Results and discussion
6.3.3 Adipogenic differentiation of 3T3-L1 preadipocytes
To evaluate their potential for soft tissue engineering, biodegradable hydrogels
(10 % initial polymer content) were seeded with 3T3-L1 preadipocytes (3-D culture,
100,000 cells per hydrogel). Two days after cell seeding, adipogenic differentiation was
induced by applying a hormonal cocktail containing insulin, corticosterone, IBMX,
and indomethacin. At day 9 after induction, the hydrogel constructs were analyzed
for their DNA content and intracellular triglyceride accumulation. For comparison,
100,000 3T3-L1 cells were seeded in 24-well plates and induced to undergo adipogenic
differentiation (conventional 2-D culture).
In 2-D culture, the seeded 3T3-L1 cells proliferated during the initial culture period.
At day 9 after induction, approx. 260,000 cells per well were found as determined
by the DNA assay (Figure 6.3C). In contrast to that, cell proliferation seemed to be
suppressed when 3T3-L1 cells were cultured within hydrogels. After the same time
period, only 82,000 cells per hydrogel were detected (Figure 6.3C). This was consistent
with the initial cell number, as determined in separate experiments in which gel samples
were seeded with 3T3-L1 cells and immediately analyzed for their DNA content (data
not shown). When regarding the intracellular triglyceride accumulation, the opposite
trend was observed (Figure 6.3D). Compared to conventional 2-D culture, the amount
of triglycerides accumulated per 100,000 cells increased approx. threefold when 3T3-L1
adipocytes were cultured in hydrogels. These findings were also illustrated by phase
contrast images. Under 2-D culture conditions, the cells were well spread and showed
a large number of small lipid droplets (Figure 6.3A). Within hydrogels, in contrast,
3T3-L1 adipocytes had a rounded morphology and contained comparatively large
vacuoles that occupied most of the cytoplasm (Figure 6.3B).
Altogether, the developed hydrogels seemed to provide a suitable environment
for 3T3-L1 preadipocytes to differentiate into mature adipocytes. Compared to
standard 2-D cell culture, the adipogenic differentiation of 3T3-L1 cells was obviously
enhanced under 3-D culture conditions. Adipogenesis has been reported to be
inhibited when preadipocytes were cultured on strongly adhesive, fibronectin coated
substrates [193, 194]. Under these culture conditions, morphological changes were
prevented and the induction of lipogenic enzymes (such as fatty acid synthetase)
was sharply reduced. However, the inhibitory effects of fibronectin on preadipocyte
119
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
*
*
2-D culture 3-D culture
0
50
100
150
200
250
300
350
Cell
nu
mb
er
(x1,0
00)
A) B)
C) D)
*
*
2-D culture 3-D culture
0
50
100
150
200
250
300
Tri
gly
ceri
des
(µg
/100,0
00
cells)
Figure 6.3: Microscopical images of 3T3-L1 cells at day 9 after induction. Adipocytes werecultured in 2-D cell culture (A) and within biodegradable hydrogels (B). Cell numberdetermined at day 9 after induction (C). Triglyceride accumulation of 3T3-L1 adipocytescultivated under 2-D and 3-D conditions (D). Data represent means ± standard deviationof three biological replicates; * indicates statistically significant differences between thetested groups (p < 0.05).
differentiation could be reversed by keeping the cells in a rounded configuration e.g.
by disrupting the cytoskeleton [193, 194]. A similar effect may explain the differences
between the tested culture conditions. In 2-D cell culture, the 3T3-L1 adipocytes
strongly adhered to the polystyrene substrate and spread over a large surface area.
In contrast to the very adhesive polystyrene, PEG-based hydrogels effectively prevent
the adsorption of ECM proteins such as fibronectin, laminin, and vitronectin [10].
Consequently, attached or embedded cells will exhibit only loose interactions with
the gel network and assume a more rounded morphology. These loose interactions
are thought to be necessary to allow the morphological changes that accompany
adipogenic differentiation [195].
On the other hand, ECM components and in particular laminin have been shown
to play a pivotal role in preadipocyte differentiation [196]. Furthermore, cell adhesion
is known to be critical for the viability of anchorage-dependent cells [177]. Designing
120
6.3 Results and discussion
suitable scaffolds for adipose tissue engineering will, therefore, require a balance be-
tween cell attachment, spreading, and differentiation [194]. Entrapping preadipocytes
into weakly adhesive hydrogel matrices may satisfy these criteria. For this purpose,
non-adhesive PEG hydrogels (degradable and non-degradable) were functionalized
with the synthetic nonapeptide Cys–Asp–Pro–Gly–Tyr–Ile–Gly–Ser–Arg (YIGSR),
which has been identified as the major cell-binding site in the B1 chain of laminin [197].
With regard to the detected cell numbers (determined at day 9 after induction),
no significant differences were detected between hydrogels containing no YIGSR and
those containing 1 µmol/mL YIGSR (Figure 6.4E). The DNA content of degradable
and non-degradable hydrogels was also comparable. Similar results have been reported
by Patel et al. for preadipocytes cultivated in photopolymerizable PEG hydrogels. Cell
proliferation was observed in biodegradable gels only after four days of cultivation [177].
In the present study, however, 3T3-L1 preadipocytes were induced to undergo adi-
pogenic differentiation before significant cell proliferation had occurred (induction
two days after cell seeding). When comparing the intracellular triglyceride accumu-
lation, significant differences were observed depending on the addition of YIGSR
(Figure 6.4F). In degradable hydrogels, the modification with YIGSR increased the
intracellular triglyceride accumulation from approx. 260 µg per 100,000 cells to approx.
400 µg per 100,000 cells. A similar trend was observed in non-degradable hydrogels.
These findings were also reflected in microscopical bright field images. Compared to
hydrogels without YIGSR (Figure 6.4A and C), 3T3-L1 preadipocytes cultured in
gels containing 1 µmol/mL YIGSR seemed to be considerably enlarged with vacuoles
that occupied most of the cytoplasm (Figure 6.4B and D). Together with the out-
come of 2-D cell culture experiments, these results clearly demonstrate the effects of
substrate stiffness and adhesiveness on adipocyte differentiation. Hydrogels function-
alized with integrin-binding motifs obviously provide the right balance between cell
attachment and differentiation, and promote the intracellular lipid accumulation of
differentiated adipocytes.
To study the effects of substrate degradability on tissue development, non-degradable
and degradable hydrogels were seeded with 3T3-L1 preadipocytes (5,000,000 cells per
hydrogel construct) and cultured over 6 weeks. At day 42 after induction, microscopical
bright field images revealed clear differences between the tested hydrogels. Within
non-degradable gels, cells were not able to form coherent tissue structures and only
121
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
A) B)
C) D)
E) F)
Degradable Non-degradable
0
20
40
60
80
100
120
Cell
nu
mb
er
(x1,0
00)
without YIGSR1 µmol/ml YIGSR
*
*
Degradable Non-degradable
0
100
200
300
400
500
600
Tri
gly
ceri
des
(µg
/100,0
00
cells)
without YIGSR1 µmol/ml YIGSR
Figure 6.4: 3T3-L1 adipocytes cultured in degradable gels without YIGSR (A), degrad-able gels with 1 µmol/mL YIGSR (B), non-degradable gels without YIGSR (C), andnon-degradable gels with 1 µmol/mL YIGSR (D). Cell number (E) and triglyceride accu-mulation (F) of 3T3-L1 cells determined at day 9 after induction. Data represent means± standard deviation of three biological replicates; * indicates statistically significantdifferences between the tested groups (p < 0.05).
isolated adipocytes were found. The individual cells contained multiple lipid droplets,
however no unilocolar fat cells were detected (Figure 6.5A and C). In contrast
to that, adipose tissue-like structures were formed when 3T3-L1 adipocytes were
cultured in enzymatically degradable gels; single cells were not detectable (Figure 6.5B
and D). Compared to non-degradable hydrogels, the intracellular lipid droplets were
considerably enlarged, resulting in many unilocular cells, a typical feature of mature
adipocytes. Altogether, these first experiments clearly indicate the advantage of
122
6.4 Conclusion
enzymatically degradable hydrogels for adipose tissue engineering. The developed gels
were sufficiently stable to withstand the manipulations associated with cell culture and
provided a suitable microenvironment for the differentiation of adipocytes. In case of
degradable hydrogels, the polymer network can be disintegrated by cell-secreted and
cell-activated proteases (such as MMPs), which obviously promotes the development
of coherent adipose tissue-like structures.
A) B)
C) D)
Figure 6.5: Phase contrast images of 3T3-L1 cells at day 42 after induction (A and B).The hydrogel constructs were stained with oil red O to visualize the size and amountof cytoplasmic lipid vesicles (C and D). 3T3-L1 adipocytes were cultured within non-degradable (A and C) and degradable hydrogels (B and D).
6.4 Conclusion
In summary, we successfully prepared PEG-based hydrogels that proved to be suitable
as 3-D scaffolds for soft tissue engineering applications. The gels were cross-linked
in the presence of cells without the use of free-radical initiators, showed mechanical
properties close to those of adipose tissue, and were susceptible to proteolytic break-
123
Chapter 6 Biointeractive hydrogels for adipose tissue engineering
down. In contrast to previously described approaches, the developed hydrogels could
be easily functionalized with adhesion peptides without requiring chemical modifica-
tions of these molecules. Cell culture experiments indicated that these biointeractive
hydrogels provide a suitable 3-D environment for 3T3-L1 cells to differentiate into
adipocytes. Long-term studies suggested that enzymatically degradable hydrogels
promote the formation of coherent tissue-like structures. In future experiments, cell
seeding and culture conditions have to be further optimized in order to generate
mature fat pads. Similar to the introduction of adhesion peptides, the hydrogels
could also be functionalized with hormones or growth factors (such as insulin, basic
fibroblast growth factor, or vascular endothelial growth factor). Since these molecules
will be covalently attached to the gel network, hormones or growth factors are expected
to interact only with encapsulated or invading cells and not with the surrounding
tissue. In future applications, the polymer-cell mixture could be supplemented with
biologically active molecules, directly injected into soft tissue defects, and cross-linked
in vivo to generate functional tissue substitutes.
124
Chapter 7
Summary and conclusions
125
Chapter 7 Summary and conclusions
Summary
This thesis was focused on the development and characterization of PEG-based
hydrogels for controlling drug delivery and promoting tissue regeneration. While
various gelation methods (i.e. ionic, hydrophobic, or covalent interactions) can be
used for the formation of injectable hydrogels [73], chemical or covalent cross-linking
is favored because it results in stable gel structures with tunable physicochemical
properties. Furthermore, physically cross-linked hydrogels (e.g. gels prepared from
PEG-b-PLGA-b-PEG triblock copolymers) often become turbid during gelation [73,
198], which excludes them from applications in ophthalmology. For these reasons,
hydrogels were synthesized by step-growth polymerization of amino-functionalized
PEG macromers with branched, amine-reactive derivatives of PEG (Figure 7.1). This
reaction could be performed under ambient conditions (pH 7.4, 37 ◦C) without the use
of free-radical initiators and allowed for the incorporation of fragile biomolecules (e.g.
proteins or nucleic acids) or living cells. The developed hydrogels can serve as inert
space-filling agents, as carrier systems for the controlled release of drug molecules, or
as three-dimensional scaffolds in tissue engineering approaches.
In first experiments, non-degradable hydrogels were synthesized that may act as in-
jectable, biologically inert substitutes for the vitreous body (Chapter 3). Transparent
hydrogels were formed by reaction between branched PEG-succinimidyl propionates
and two different types of PEG-amines (Figure 7.1 and 7.2). The gels were char-
acterized by oscillatory rheometry and 1H-NMR experiments. The liquid precursor
solutions easily passed through small-gauge needles but solidified within 5 to 10 min,
which would be appropriate for intraocular injections. By varying the concentration
of macromers, the functionality of the PEG-amine, and the conditions during cross-
linking, gels with mechanical properties similar to those of the natural vitreous body
were obtained. The cross-linked hydrogels showed no cytotoxic effects and may be
used as vitreous substitutes or as intraocular drug delivery systems.
As shown in Chapter 3, the developed hydrogels have great promise as matrices
for the controlled release of macromolecules. To enable the efficient characterization of
hydrogel-based drug delivery systems, mechanical testing, fluorescence recovery after
photobleaching (FRAP), and pulsed field gradient NMR spectroscopy were investigated
as alternatives to release experiments (Chapter 4). The developed hydrogels were
126
OO
OO
n*
O
N
O
O
OO
OO
n*
O
N
O
O
OO
OH
2N
n*
OO
OH
2N
n*
OO
ON H
O
O
O
ON
O
O
n*
OO
ON H
n*
H NN H
O
O
O
N
O
H2NH3C
Co
mp
on
en
tA
Hy
dro
ge
lC
om
po
ne
nt
B
4arm
PE
G10k-d
Tyr-
SC
4arm
PE
G10k-S
PA
4arm
PE
G10k-S
PA
4arm
PE
G10k-N
H2
4arm
PE
G10k-L
GP
A
4arm
PE
G10k-N
H2
hyd
roly
tically
deg
rad
ab
le
no
n-d
eg
rad
ab
le
en
zym
ati
cally
deg
rad
ab
le
Fig
ure
7.1:
Pol
ymer
sde
rive
dfr
omfo
ur-a
rmed
poly
(eth
ylen
egl
ycol
)(m
olec
ular
wei
ght
10kD
a,4a
rmP
EG
10k-
OH
).N
on-d
egra
dabl
ehy
drog
els
wer
epr
epar
edby
step
-gro
wth
poly
mer
izat
ion
ofbr
anch
edP
EG
-suc
cini
mid
ylpr
opio
nate
s(4
arm
PE
G10
k-SP
A)
and
bran
ched
PE
G-a
min
es(4
arm
PE
G10
k-N
H2).
Rea
ctio
nof
arom
atic
PE
G-s
ucci
nim
idyl
carb
onat
es(4
arm
PE
G10
k-dT
yr-S
C)
wit
h4a
rmP
EG
10k-
NH
2re
sult
edin
hydr
olyt
ical
lyde
grad
able
gels
.Fo
rth
epr
epar
atio
nof
enzy
mat
ical
lyde
grad
able
hydr
ogel
s,br
anch
edP
EG
-am
ines
wer
efu
ncti
onal
ized
wit
hth
eco
llage
nase
-sen
itiv
epe
ptid
eA
la–
Pro
–Gly↓L
eu(4
arm
PE
G10
k-L
GPA
)an
dcr
oss-
linke
dw
ith
4arm
PE
G10
k-SP
A.S
truc
tura
lcha
nges
ofth
eP
EG
mac
rom
ers
(e.g
.va
riat
ions
inth
em
olec
ular
wei
ght
and/
ornu
mbe
rof
bran
ches
)w
ould
addi
tion
ally
alte
rth
ege
lpr
oper
ties
.
127
Chapter 7 Summary and conclusions
Figure 7.2: PEG hydrogel formed by reaction between branched PEG-succinimidyl propi-onates and branched PEG-amines. The gels have a high water content (up to 97 % inthe swollen state) and are optically transparent.
loaded with FITC-dextrans and characterized for their mechanical properties and
swelling characteristics. Subsequently, the translational diffusion coefficients (D) of
the incorporated FITC-dextrans were measured. Since the determined values of D
agreed very well with those obtained from release studies, mechanical testing, FRAP,
and pulsed field gradient NMR spectroscopy can be used to rapidly evaluate the
potential of newly developed drug delivery systems.
One drawback of hydrogel-based drug delivery systems is their limited capacity
to restrict the mobility of encapsulated molecules, which usually results in relatively
rapid drug release (Chapter 3 and 4). To prolong the release rates of hydrogels, drug
molecules were covalently tethered to the gel network via hydrolytically degradable
anchor groups. For this purpose, branched PEG was functionalized with aromatic
succinimidyl carbonate groups (Figure 7.1) that readily react with amino groups of
other polymers, peptides, or proteins under formation of biodegradable carbamate
bonds (Chapter 5). The strength of the prepared hydrogels ranged from 1075 to
2435 Pa; the degradation time varied between 24 and 120 h. FRAP experiments
showed that FITC-BSA was successfully bound to the gel network. During polymer
degradation, the mobility of the tethered molecules gradually increased. Using FITC-
BSA and lysozyme as model proteins, the suitability of the developed hydrogels for
the time-controlled release of proteins was shown. The obtained release profiles had a
sigmoidal shape; protein release and gel degradation occurred simultaneously.
In the last study, the virtually inert PEG hydrogels were modified to become
biointeractive and used as three-dimensional scaffolds for adipose tissue engineering.
Since substrate mechanics and degradability were recognized to have a strong influence
on cell differentiation and tissue morphogenesis (Chapter 2), branched PEG-amines
were modified with the collagenase-sensitive peptide sequence Ala–Pro–Gly↓Leu. To
128
form hydrogels, these peptide-modified PEG-amines were cross-linked with branched
PEG-succinimidyl propionates (Figure 7.1). The gel strength ranged from 1708 to
7412 Pa, depending on the initial polymer concentration. To mediate cell adhesion,
the gels were functionalized with the integrin-binding motif Tyr–Ile–Gly–Ser–Arg
(YIGSR). These hydrogels mimic the cellular recognition of the natural extracellular
matrix (ECM) and were degraded by cell-secreted proteases. Cell culture experiments
clearly demonstrated the suitability of these biointeractive hydrogels for soft tissue
regeneration (Chapter 6). Compared to standard two-dimensional cell culture, the
developed hydrogels significantly enhanced the intracellular triglyceride accumulation
of encapsulated 3T3-L1 adipocytes. Functionalization with YIGSR further enhanced
lipid synthesis of differentiating adipocytes. Long-term studies suggested that en-
zymatically degradable hydrogels furthermore promote the formation of coherent
adipose tissue-like structures.
Conclusions and outlook
In conclusion, branched PEG proved to be the ideal starting material for the synthesis
of hydrogels for regenerative medicine. Using a combinatorial approach, a variety of
hydrogels could be prepared from comparatively few building blocks. The synthesized
macromers allowed for the preparation of non-degradable, hydrolytically degradable,
and enzymatically degradable hydrogels (Figure 7.1). Cross-linking could be performed
in the presence of cells without the use of free-radical initiators. In a recent study
beyond this thesis, non-degradable hydrogels were injected into enucleated porcine
eyes [199]. These gels solidified approx. 5 min after injection and remained optically
transparent. The tissue compatibility of the developed hydrogels was evaluated in vitro
using a perfusion organ culture model of full-thickness porcine retina (Figure 7.3) [200,
201]. Altogether, these non-degradable hydrogels have great promise as long-term
substitutes for the natural vitreous body [199, 200].
Because of their amine-reactive character, the gel-forming polymers can also be
functionalized with biologically active peptides or proteins (e.g. adhesion peptides,
growth factors, or antibodies) without requiring chemical modifications of these
molecules. In future applications, therapeutic peptides or proteins could be dissolved
129
Chapter 7 Summary and conclusions
A B C
100 µm
Figure 7.3: PEG hydrogels in perfusion organ culture of adult porcine retina (A) and afterfour days of perfusion (B). Light micrograph of porcine retina stained with haematoxylinand eosin (C). Histologically, the retinal structure remained well preserved [200]. Scalebar represents 100 µm.
together with the gel-forming polymers and injected into the patient in a minimally
invasive manner (e.g. into the vitreous cavity). During gelation, these molecules are
covalently tethered to the gel network, which effectively prevents their immediate
release. If hydrolytically degradable polymers are used for gel preparation, this will
allow for the time-controlled release of therapeutic peptide or proteins. Potential
candidates for pharmacologically active proteins include antibodies against vascular
endothelial growth factor (VEGF), a growth factor that is involved in the pathogenesis
of proliferative diabetic retinopathy (PDR).
For applications in soft tissue engineering, enzymatically degradable hydrogels
seem to be especially promising. These gels are sufficiently stable to withstand the
manipulations associated with implantation and provide a suitable microenvironment
that directs cell proliferation and differentiation. Once placed at the application site,
the gels are degraded by cell-secreted proteases; matrix degradation is expected to
occur in spatial and temporal synchrony with the deposition of ECM. As described
above, the hydrogels can be easily functionalized with integrin-binding peptides or
growth factors. This creates biointeractive hydrogels that mimic the complexity of
the natural ECM. Structural changes of the PEG macromers (e.g. variations in the
molecular weight and/or number of branches) would provide additional control over
the mechanical properties, swelling behavior, and degradation rate of the formed
hydrogels. Although further experiments will have to be carried out in the future,
these first studies demonstrate that the developed hydrogels have great promise for a
number of applications in regenerative medicine.
130
Bibliography
[1] R. Langer and J. P. Vacanti. Tissue engineering. Science, 260(5110):920–926,
1993.
[2] S. Petit-Zeman. Regenerative medicine. Nat. Biotechnol., 19(3):201–206, 2001.
[3] K. Ghosh and D. E. Ingber. Micromechanical control of cell and tissue develop-
ment: Implications for tissue engineering. Adv. Drug Deliv. Rev., 59(13):1306–
1318, 2007.
[4] A. Oosterlee and A. Rahmel, editors. Annual Report 2008. Eurotransplant
International Foundation, Leiden, 2008.
[5] R. R. Chen and D. J. Mooney. Polymeric growth factor delivery strategies for
tissue engineering. Pharm. Res., 20(8):1103–1112, 2003.
[6] L. Cao and D. J. Mooney. Spatiotemporal control over growth factor signaling
for therapeutic neovascularization. Adv. Drug Deliv. Rev., 59(13):1340 –1350,
2007.
[7] J. K. Teßmar and A. M. Gopferich. Matrices and scaffolds for protein delivery
in tissue engineering. Adv. Drug Deliv. Rev., 59(4-5):274–291, 2007.
[8] M. P. Lutolf and J. A. Hubbell. Synthetic biomaterials as instructive extracellular
microenvironments for morphogenesis in tissue engineering. Nat. Biotechnol.,
23(1):47–55, 2005.
131
Bibliography
[9] A. S. Hoffman. Hydrogels for biomedical applications. Adv. Drug Deliv. Rev.,
54(1):3–12, 2002.
[10] J. L. Drury and D. J. Mooney. Hydrogels for tissue engineering: scaffold design
variables and applications. Biomaterials, 24(24):4337–4351, 2003.
[11] C.-C. Lin and K. Anseth. PEG hydrogels for the controlled release of
biomolecules in regenerative medicine. Pharm. Res., 26(3):631–643, 2009.
[12] T. R. Hoare and D. S. Kohane. Hydrogels in drug delivery: Progress and
challenges. Polymer, 49(8):1993–2007, 2008.
[13] J. Teßmar, F. Brandl, and A. Gopferich. Fundamentals of tissue engineering and
regenerative medicine, chapter Hydrogels for tissue engineering, pages 495–518.
Springer, Berlin, 2009.
[14] J. Harris. Poly(ethylene glycol) chemistry: biotechnical and biomedical applica-
tions. Plenum Press, New York, 1992.
[15] H. Shin, S. Jo, and A. G. Mikos. Biomimetic materials for tissue engineering.
Biomaterials, 24(24):4353–4364, 2003.
[16] J. A. Hubbell. Materials as morphogenetic guides in tissue engineering. Curr.
Opin. Biotechnol., 14(5):551–558, 2003.
[17] E. A. Silva and D. J. Mooney. Synthetic extracellular matrices for tissue
engineering and regeneration. Curr. Top. Dev. Biol., 64:181–205, 2004.
[18] I. Kocur and S. Resnikoff. Visual impairment and blindness in Europe and their
prevention. Br. J. Ophthalmol., 86(7):716–722, 2002.
[19] U. S. Schwarz and I. B. Bischofs. Physical determinants of cell organization in
soft media. Med. Eng. Phys., 27(9):763–772, 2005.
[20] J. Y. Wong, J. B. Leach, and X. Q. Brown. Balance of chemistry, topography, and
mechanics at the cell-biomaterial interface: Issues and challenges for assessing
the role of substrate mechanics on cell response. Surf. Sci., 570(1-2):119–133,
2004.
132
Bibliography
[21] D. E. Discher, P. Janmey, and Y.-l. Wang. Tissue cells feel and respond to the
stiffness of their substrate. Science, 310(5751):1139–1143, 2005.
[22] A. Curtis and M. Riehle. Tissue engineering: the biophysical background. Phys.
Med. Biol., 46(4):R47–R65, 2001.
[23] L. G. Griffith. Emerging design principles in biomaterials and scaffolds for tissue
engineering. Ann. N. Y. Acad. Sci., 961(1):83–95, 2002.
[24] W. F. Liu and C. S. Chen. Engineering biomaterials to control cell function.
Mater. Today, 8(12):28–35, 2005.
[25] K. Y. Lee and D. J. Mooney. Hydrogels for tissue engineering. Chem. Rev.,
101(7):1869–1880, 2001.
[26] J. Vitte, A. M. Benoliel, A. Pierres, and P. Bongrand. Is there a predictable
relationship between surface physical-chemical properties and cell behaviour at
the interface? Eur. Cell Mater., 7:52–63, 2004.
[27] Y. Tabata. Tissue regeneration based on growth factor release. Tissue Eng.,
9(Suppl. 1):S5–S15, 2003.
[28] C. Sfeir, J. Jadlowiec, H. Koch, and P. Campbell. Signaling molecules for tissue
engineering. In J. Hollinger, T. Einhorn, B. Doll, and C. Sfeir, editors, Bone
Tissue Engineering, pages 125–147. CRC Press, Boca Raton, 2005.
[29] R. Vasita and D. S. Katti. Growth factor-delivery systems for tissue engineering:
a materials perspective. Expert Rev. Med. Devices, 3(1):29–47, 2006.
[30] S. B. Carter. Principles of cell motility: the direction of cell movement and
cancer invasion. Nature, 208(16):1183–1187, 1965.
[31] S. B. Carter. Haptotaxis and the mechanism of cell motility. Nature, 213(73):256–
260, 1967.
[32] B. K. Brandley and R. L. Schnaar. Tumor cell haptotaxis on covalently immo-
bilized linear and exponential gradients of a cell adhesion peptide. Dev. Biol.,
135(1):74–86, 1989.
133
Bibliography
[33] M. E. Chicurel, C. S. Chen, and D. E. Ingber. Cellular control lies in the balance
of forces. Curr. Opin. Cell Biol., 10(2):232–239, 1998.
[34] D. E. Ingber. Tensegrity I. Cell structure and hierarchical systems biology. J.
Cell Sci., 116(7):1157–1173, 2003.
[35] D. E. Ingber. Tensegrity II. How structural networks influence cellular informa-
tion processing networks. J. Cell Sci., 116(8):1397–1408, 2003.
[36] A. J. Garcia. Get a grip: integrins in cell-biomaterial interactions. Biomaterials,
26(36):7525–7529, 2005.
[37] B. Geiger, A. Bershadsky, R. Pankov, and K. M. Yamada. Transmembrane
crosstalk between the extracellular matrix and the cytoskeleton. Nat. Rev. Mol.
Cell Biol., 2(11):793–805, 2001.
[38] B. Geiger and A. Bershadsky. Exploring the neighborhood: adhesion-coupled
cell mechanosensors. Cell, 110(2):139–142, 2002.
[39] E. Cukierman, R. Pankov, D. R. Stevens, and K. M. Yamada. Taking cell-matrix
adhesions to the third dimension. Science, 294(5547):1708–1712, 2001.
[40] E. Cukierman, R. Pankov, and K. M. Yamada. Cell interactions with three-
dimensional matrices. Curr. Opin. Cell Biol., 14(5):633–640, 2002.
[41] D. Choquet, D. P. Felsenfeld, and M. P. Sheetz. Extracellular matrix rigidity
causes strengthening of integrin-cytoskeleton linkages. Cell, 88(1):39–48, 1997.
[42] C. M. Lo, H. B. Wang, M. Dembo, and Y. L. Wang. Cell movement is guided
by the rigidity of the substrate. Biophys. J., 79(1):144–152, 2000.
[43] D. S. Gray, J. Tien, and C. S. Chen. Repositioning of cells by mechanotaxis
on surfaces with micropatterned Young’s modulus. J. Biomed. Mater. Res. A,
66A(3):605–614, 2003.
[44] R. J. Pelham and Y.-l. Wang. Cell locomotion and focal adhesions are regulated
by substrate flexibility. Proc. Natl. Acad. Sci. U.S.A., 94(25):13661–13665,
1997.
134
Bibliography
[45] H. B. Wang, M. Dembo, and Y.-l. Wang. Substrate flexibility regulates growth
and apoptosis of normal but not transformed cells. Am. J. Physiol. Cell Physiol.,
279(5):C1345–C1350, 2000.
[46] B. Vailhe, X. Ronot, P. Tracqui, Y. Usson, and L. Tranqui. In vitro angiogenesis
is modulated by the mechanical properties of fibrin gels and is related to αvβ3
integrin localization. In Vitro Cell. Dev. Biol. Anim., 33(10):763–773, 1997.
[47] C. F. Deroanne, C. M. Lapiere, and B. V. Nusgens. In vitro tubulogenesis of
endothelial cells by relaxation of the coupling extracellular matrix-cytoskeleton.
Cardiovasc. Res., 49(3):647–658, 2001.
[48] L. A. Flanagan, Y. E. Ju, B. Marg, M. Osterfield, and P. A. Janmey. Neurite
branching on deformable substrates. Neuroreport, 13(18):2411–2415, 2002.
[49] A. D. Bershadsky, N. Q. Balaban, and B. Geiger. Adhesion-dependent cell
mechanosensitivity. Annu. Rev. Cell Dev. Biol., 19(1):677–695, 2003.
[50] J. A. Rowley and D. J. Mooney. Alginate type and RGD density control
myoblast phenotype. J. Biomed. Mater. Res., 60(2):217–223, 2002.
[51] A. Engler, L. Bacakova, C. Newman, A. Hategan, M. Griffin, and D. Discher.
Substrate compliance versus ligand density in cell on gel responses. Biophys. J.,
86(1):617–628, 2004.
[52] S. R. Peyton and A. J. Putnam. Extracellular matrix rigidity governs smooth
muscle cell motility in a biphasic fashion. J. Cell. Physiol., 204(1):198–209,
2005.
[53] H. Abe, K. Hayashi, and M. Sato, editors. Data Book on Mechanical Properties
of Living Cells, Tissues, and Organs. Springer-Verlag, Tokyo, 1996.
[54] K. S. Anseth, C. N. Bowman, and L. Brannon-Peppas. Mechanical properties of
hydrogels and their experimental determination. Biomaterials, 17(17):1647–1657,
1996.
135
Bibliography
[55] R. K. Korhonen, M. S. Laasanen, J. Toyras, J. Rieppo, J. Hirvonen, H. J.
Helminen, and J. S. Jurvelin. Comparison of the equilibrium response of articular
cartilage in unconfined compression, confined compression and indentation. J.
Biomech., 35(7):903–909, 2002.
[56] H. A. Barnes, J. F. Hutton, and K. Walters, editors. An introduction to rheology.
Elsevier Science Publishers B.V., Amsterdam, 1989.
[57] K. Ichihara, T. Taguchi, I. Sakuramoto, S. Kawano, and S. Kawai. Mechanism
of the spinal cord injury and the cervical spondylotic myelopathy: new approach
based on the mechanical features of the spinal cord white and gray matter. J
Neurosurg., 99(3 Suppl):278–285, 2003.
[58] K. A. Athanasiou, A. Agarwal, and F. J. Dzida. Comparative study of the
intrinsic mechanical properties of the human acetabular and femoral head
cartilage. J. Orthop. Res., 12(3):340–349, 1994.
[59] D. Raftopoulos, E. Katsamanis, F. Saul, W. Liu, and S. Saddemi. An interme-
diate loading rate technique for the determination of mechanical properties of
human femoral cortical bone. J. Biomed. Eng., 15(1):60–66, 1993.
[60] J. Domke and M. Radmacher. Measuring the elastic properties of thin polymer
films with the atomic force microscope. Langmuir, 14(12):3320–3325, 1998.
[61] W. R. Bowen, R. W. Lovitt, and C. J. Wright. Application of atomic force
microscopy to the study of micromechanical properties of biological materials.
Biotechnol. Lett., 22(11):893–903, 2000.
[62] A. J. Engler, L. Richert, J. Y. Wong, C. Picart, and D. E. Discher. Surface probe
measurements of the elasticity of sectioned tissue, thin gels and polyelectrolyte
multilayer films: correlations between substrate stiffness and cell adhesion. Surf.
Sci., 570(1-2):142–154, 2004.
[63] A. Manduca, T. E. Oliphant, M. A. Dresner, J. L. Mahowald, S. A. Kruse,
E. Amromin, J. P. Felmlee, J. F. Greenleaf, and R. L. Ehman. Magnetic
resonance elastography: non-invasive mapping of tissue elasticity. Med. Image
Anal., 5(4):237–254, 2001.
136
Bibliography
[64] G. Madelin, N. Baril, J. D. De Certaines, J.-M. Franconi, and E. Thiaudiere.
NMR characterization of mechanical waves. In G. Webb, editor, Annual Reports
on NMR Spectroscopy, volume Volume 53, pages 203–244. Academic Press,
2004.
[65] S. I. Ringleb, Q. Chen, D. S. Lake, A. Manduca, R. L. Ehman, and K. N. An.
Quantitative shear wave magnetic resonance elastography: comparison to a
dynamic shear material test. Magn. Reson. Med., 53(5):1197–1201, 2005.
[66] S. F. Othman, H. Xu, T. J. Royston, and R. L. Magin. Microscopic magnetic
resonance elastography (µMRE). Magn. Reson. Med., 54(3):605–615, 2005.
[67] H. J. Kong, T. R. Polte, E. Alsberg, and D. J. Mooney. FRET measurements
of cell-traction forces and nano-scale clustering of adhesion ligands varied by
substrate stiffness. Proc. Natl. Acad. Sci. U.S.A., 102(12):4300–4305, 2005.
[68] J. Szollosi, S. Damjanovich, and L. Matyus. Application of fluorescence resonance
energy transfer in the clinical laboratory: routine and research. Cytometry,
34(4):159–179, 1998.
[69] E. A. Jares-Erijman and T. M. Jovin. FRET imaging. Nat. Biotechnol.,
21(11):1387–1395, 2003.
[70] A. Baruch, D. A. Jeffery, and M. Bogyo. Enzyme activity – it’s all about image.
Trends Cell Biol., 14(1):29–35, 2004.
[71] O. Smidsrød and G. Skjak-Bræk. Alginate as immobilization matrix for cells.
Trends Biotechnol., 8:71–78, 1990.
[72] A. S. Sawhney, C. P. Pathak, and J. A. Hubbell. Bioerodible hydrogels based
on photopolymerized poly(ethylne glycol)-co-poly(α-hydroxy acid) diacrylate
macromers. Macromolecules, 26(4):581–587, 1993.
[73] W. E. Hennink and C. F. van Nostrum. Novel crosslinking methods to design
hydrogels. Adv. Drug Deliv. Rev., 54(1):13–36, 2002.
137
Bibliography
[74] M. P. Lutolf and J. A. Hubbell. Synthesis and physicochemical characterization
of end-linked poly(ethylene glycol)-co-peptide hydrogels formed by Michael-type
addition. Biomacromolecules, 4(3):713–722, 2003.
[75] E. Ruel-Gariepy and J. C. Leroux. In situ-forming hydrogels–review of
temperature-sensitive systems. Eur. J. Pharm. Biopharm., 58(2):409–426,
2004.
[76] S. Zhang. Fabrication of novel biomaterials through molecular self-assembly.
Nat. Biotechnol., 21(10):1171–1178, 2003.
[77] K. Y. Lee, J. A. Rowley, P. Eiselt, E. M. Moy, K. H. Bouhadir, and D. J.
Mooney. Controlling mechanical and swelling properties of alginate hydrogels
independently by cross-linker type and cross-linking density. Macromolecules,
33(11):4291–4294, 2000.
[78] M. A. LeRoux, F. Guilak, and L. A. Setton. Compressive and shear properties
of alginate gel: effects of sodium ions and alginate concentration. J. Biomed.
Mater. Res., 47(1):46–53, 1999.
[79] J. W. Gunn, S. D. Turner, and B. K. Mann. Adhesive and mechanical properties
of hydrogels influence neurite extension. J. Biomed. Mater. Res. A, 72A(1):91–97,
2005.
[80] S. J. Bryant and K. S. Anseth. Hydrogel properties influence ECM production by
chondrocytes photoencapsulated in poly(ethylene glycol) hydrogels. J. Biomed.
Mater. Res., 59(1):63–72, 2002.
[81] G. M. Cruise, D. S. Scharp, and J. A. Hubbell. Characterization of permeability
and network structure of interfacially photopolymerized poly(ethylene glycol)
diacrylate hydrogels. Biomaterials, 19(14):1287–1294, 1998.
[82] Y. Park, M. P. Lutolf, J. A. Hubbell, E. B. Hunziker, and M. Wong. Bovine
primary chondrocyte culture in synthetic matrix metalloproteinase-sensitive
poly(ethylene glycol)-based hydrogels as a scaffold for cartilage repair. Tissue
Eng., 10(3-4):515–522, 2004.
138
Bibliography
[83] R. H. Li, D. H. Altreuter, and F. T. Gentile. Transport characterization of
hydrogel matrices for cell encapsulation. Biotechnol. Bioeng., 50(4):365–373,
1996.
[84] H. A. Leddy, H. A. Awad, and F. Guilak. Molecular diffusion in tissue-engineered
cartilage constructs: effects of scaffold material, time, and culture conditions. J.
Biomed. Mater. Res. B Appl. Biomater., 70(2):397–406, 2004.
[85] S. P. Baldwin and M. W. Saltzman. Materials for protein delivery in tissue
engineering. Adv Drug Deliver Rev, 33(1-2):71–86, 1998.
[86] K. Kellner, G. Liebsch, I. Klimant, O. S. Wolfbeis, T. Blunk, M. B. Schulz, and
A. Gopferich. Determination of oxygen gradients in engineered tissue using a
fluorescent sensor. Biotechnol. Bioeng., 80(1):73–83, 2002.
[87] E. Alsberg, H. Kong, Y. Hirano, M. K. Smith, A. Albeiruti, and D. J. Mooney.
Regulating bone formation via controlled scaffold degradation. J. Dent. Res.,
82(11):903–908, 2003.
[88] H. J. Kong, D. Kaigler, K. Kim, and D. J. Mooney. Controlling rigidity and
degradation of alginate hydrogels via molecular weight distribution. Biomacro-
molecules, 5(5):1720–1727, 2004.
[89] T. Boontheekul, H.-J. Kong, and D. J. Mooney. Controlling alginate gel degra-
dation utilizing partial oxidation and bimodal molecular weight distribution.
Biomaterials, 26(15):2455–2465, 2005.
[90] M. P. Lutolf, J. L. Lauer-Fields, H. G. Schmoekel, A. T. Metters, F. E. Weber,
G. B. Fields, and J. A. Hubbell. Synthetic matrix metalloproteinase-sensitive
hydrogels for the conduction of tissue regeneration: engineering cell-invasion
characteristics. Proc. Natl. Acad. Sci. U.S.A., 100(9):5413–5418, 2003.
[91] K. J. Gooch and C. J. Tennant, editors. Mechanical forces: their effects on cells
and tissues. Springer-Verlag, Berlin, 1997.
[92] A. Katsumi, A. W. Orr, E. Tzima, and M. A. Schwartz. Integrins in mechan-
otransduction. J. Biol. Chem., 279(13):12001–12004, 2004.
139
Bibliography
[93] G. P. Dillon, X. Yu, A. Sridharan, J. P. Ranieri, and R. V. Bellamkonda. The
influence of physical structure and charge on neurite extension in a 3D hydrogel
scaffold. J. Biomater. Sci. Polym. Ed., 9(10):1049–1069, 1998.
[94] A. P. Balgude, X. Yu, A. Szymanski, and R. V. Bellamkonda. Agarose gel
stiffness determines rate of DRG neurite extension in 3D cultures. Biomaterials,
22(10):1077–1084, 2001.
[95] M. Wong, M. Siegrist, X. Wang, and E. Hunziker. Development of mechanically
stable alginate/chondrocyte constructs: effects of guluronic acid content and
matrix synthesis. J. Orthop. Res., 19(3):493–499, 2001.
[96] A. L. Sieminski, R. P. Hebbel, and K. J. Gooch. The relative magnitudes of en-
dothelial force generation and matrix stiffness modulate capillary morphogenesis
in vitro. Exp. Cell Res., 297(2):574–584, 2004.
[97] S. J. Bryant and K. S. Anseth. Controlling the spatial distribution of ECM
components in degradable PEG hydrogels for tissue engineering cartilage. J.
Biomed. Mater. Res. A, 64(1):70–79, 2003.
[98] M. J. Mahoney and K. S. Anseth. Three-dimensional growth and function
of neural tissue in degradable polyethylene glycol hydrogels. Biomaterials,
27(10):2265–2274, 2006.
[99] L. Meinel, S. Hofmann, V. Karageorgiou, L. Zichner, R. Langer, D. Kaplan, and
G. Vunjak-Novakovic. Engineering cartilage-like tissue using human mesenchy-
mal stem cells and silk protein scaffolds. Biotechnol. Bioeng., 88(3):379–391,
2004.
[100] M. P. Lutolf, F. E. Weber, H. G. Schmoekel, J. C. Schense, T. Kohler, R. Muller,
and J. A. Hubbell. Repair of bone defects using synthetic mimetics of collagenous
extracellular matrices. Nat. Biotechnol., 21(5):513–518, 2003.
[101] M. H. Zaman, R. D. Kamm, P. Matsudaira, and D. A. Lauffenburger. Com-
putational model for cell migration in three-dimensional matrices. Biophys. J.,
89(2):1389–1397, 2005.
140
Bibliography
[102] F. Brandl, F. Sommer, and A. Gopferich. Rational design of hydrogels for
tissue engineering: Impact of physical factors on cell behavior. Biomaterials,
28(2):134–146, 2007.
[103] M. C. Cushing and K. S. Anseth. Hydrogel cell cultures. Science, 316(5828):1133–
1134, 2007.
[104] P. N. Bishop. Structural macromolecules and supramolecular organisation of
the vitreous gel. Prog. Ret. Eye Res., 19(3):323–344, 2000.
[105] N. Soman and R. Banerjee. Artificial vitreous replacements. Biomed. Mater.
Eng., 13(1):59–74, 2003.
[106] M. J. Colthurst, R. L. Williams, P. S. Hiscott, and I. Grierson. Biomaterials
used in the posterior segment of the eye. Biomaterials, 21(7):649–665, 2000.
[107] T. V. Chirila, Y. Hong, P. D. Dalton, I. J. Constable, and M. F. Refojo. The use
of hydrophilic polymers as artificial vitreous. Prog. Polym. Sci., 23(3):475–508,
1998.
[108] P. Gargiulo, C. Giusti, D. Pietrobono, D. La Torre, D. Diacono, and G. Tambur-
rano. Diabetes mellitus and retinopathy. Dig. Liver Dis., 36(Supplement 1):S101–
S105, 2004.
[109] P. D. Dalton, T. V. Chirila, Y. Hong, and A. Jefferson. Oscillatory shear
experiments as criteria for potential vitreous substitutes. Polym. Gels Netw.,
3(4):429–444, 1995.
[110] T. Yasukawa, Y. Ogura, Y. Tabata, H. Kimura, P. Wiedemann, and Y. Honda.
Drug delivery systems for vitreoretinal diseases. Prog. Ret. Eye Res., 23(3):253–
281, 2004.
[111] W. Eichler, Y. Yafai, P. Wiedemann, and D. Fengler. Antineovascular agents in
the treatment of eye diseases. Curr. Pharm. Des., 12(21):2645–2660, 2006.
[112] C. A. P. Quinn, R. E. Connor, and A. Heller. Biocompatible, glucose-permeable
hydrogel for in situ coating of implantable biosensors. Biomaterials, 18(24):1665–
1670, 1997.
141
Bibliography
[113] G. M. C. Wallace. A tissue sealant based on reactive multifunctional polyethylene
glycol. J. Biomed. Mater. Res., 58(5):545–555, 2001.
[114] M. Wathier, M. S. Johnson, M. A. Carnahan, C. Baer, B. W. McCuen, and M. W.
Kim, T.; Grinstaff. In situ polymerized hydrogels for repairing scleral incisions
used in pars plana vitrectomy procedures. ChemMedChem, 1(8):821–825, 2006.
[115] P. Mongondry, C. Bonnans-Plaisance, M. Jean, and J.-F. Tassin. Mild synthesis
of amino-poly(ethylene glycol)s. Application to steric stabilization of clays.
Macromol. Rapid Commun., 24(11):681–685, 2003.
[116] M. Sedaghat-Herati, P. Miller, A. Kozlowski, and J. M. Harris. Synthesis of
methoxypoly(oxyethylene)propionic acid. Polym. Bull., 43(1):35–41, 1999.
[117] A. Metters and J. Hubbell. Network formation and degradation behavior
of hydrogels formed by Michael-type addition reactions. Biomacromolecules,
6(1):290–301, 2005.
[118] M. Mensitieri, L. Ambrosio, L. Nicolais, L. Balzano, and D. Lepore. The rheo-
logical behaviour of animal vitreus and its comparison with vitreal substitutes.
J. Mater. Sci. Mater. Med., 5(9 - 10):743–747, 1994.
[119] C. S. Nickerson, H. L. Karageozian, J. Park, and J. A. Kornfield. Internal
tension: a novel hypothesis concerning the mechanical properties of the vitreous
humor. Macromol. Symp., 227(1):183–189, 2005.
[120] A. Pizzoferrato, G. Ciapetti, S. Stea, E. Cenni, C. Arciola, D. Granchi, and
L. Savarino. Cell culture methods for testing biocompatibility. Clin. Mater.,
15(3):173–190, 1994.
[121] W. W. Minuth, S. Kloth, J. Aigner, M. Sittinger, and W. Rockl. Organo-typical
environment for cultured cells and tissues. Bioforum, 17:412–416, 1994.
[122] W. A. Herrmann, K. Kobuch, S. Kloth, C. Framme, and J. Roider. Full-thickness
adult retina in perfusion culture. A new organotypical model for toxicity tests.
Invest. Ophthalmol. Vis. Sci., 40 (Suppl.):S989–, 1999.
142
Bibliography
[123] A. A. Moshfeghi and G. A. Peyman. Micro- and nanoparticulates. Adv. Drug
Deliv. Rev., 57(14):2047–2052, 2005.
[124] S. Drotleff, U. Lungwitz, M. Breunig, A. Dennis, T. Blunk, J. Teßmar, and
A. Gopferich. Biomimetic polymers in pharmaceutical and biomedical sciences.
Eur. J. Pharm. Biopharm., 58(2):385–407, 2004.
[125] F. Sommer, K. Kobuch, F. Brandl, B. Wild, C. Framme, B. Weiser, J. Teßmar,
V. P. Gabel, T. Blunk, and A. Gopferich. Ascorbic acid modulates proliferation
and extracellular matrix accumulation of hyalocytes. Tissue Eng., 13(6):1281–
1289, 2007.
[126] C.-C. Lin and A. T. Metters. Hydrogels in controlled release formulations:
network design and mathematical modeling. Adv. Drug Deliv. Rev., 58(12-
13):1379–1408, 2006.
[127] J. Siepmann and A. Gopferich. Mathematical modeling of bioerodible, polymeric
drug delivery systems. Adv. Drug Deliv. Rev., 48(2-3):229–247, 2001.
[128] J. Siepmann and F. Siepmann. Mathematical modeling of drug delivery. Int. J.
Pharm., 364(2):328–343, 2008.
[129] D. Axelrod, D. E. Koppel, J. Schlessinger, E. Elson, and W. W. Webb. Mobil-
ity measurement by analysis of fluorescence photobleaching recovery kinetics.
Biophys. J., 16(9):1055–1069, 1976.
[130] K. Braeckmans, L. Peeters, N. N. Sanders, S. C. De Smedt, and J. Demeester.
Three-dimensional fluorescence recovery after photobleaching with the confocal
scanning laser microscope. Biophys. J., 85(4):2240–2252, 2003.
[131] T. K. Meyvis, S. C. De Smedt, P. Van Oostveldt, and J. Demeester. Fluorescence
recovery after photobleaching: a versatile tool for mobility and interaction
measurements in pharmaceutical research. Pharm. Res., 16(8):1153–1162, 1999.
[132] M. C. Branco, D. J. Pochan, N. J. Wagner, and J. P. Schneider. Macro-
molecular diffusion and release from self-assembled β-hairpin peptide hydrogels.
Biomaterials, 30(7):1339–1347, 2009.
143
Bibliography
[133] M. D. Burke, J. O. Park, M. Srinivasarao, and S. A. Khan. A novel enzymatic
technique for limiting drug mobility in a hydrogel matrix. J. Controlled Release,
104(1):141–153, 2005.
[134] A. J. Kuijpers, G. H. M. Engbers, T. K. L. Meyvis, S. S. C. De Smedt, J. De-
meester, J. Krijgsveld, S. A. J. Zaat, J. Dankert, and J. Feijen. Combined
gelatin-chondroitin sulfate hydrogels for controlled release of cationic antibacte-
rial proteins. Macromolecules, 33(10):3705–3713, 2000.
[135] S. R. Van Tomme, B. G. De Geest, K. Braeckmans, S. C. De Smedt, F. Siepmann,
J. Siepmann, C. F. Van Nostrum, and W. E. Hennink. Mobility of model proteins
in hydrogels composed of oppositely charged dextran microspheres studied by
protein release and fluorescence recovery after photobleaching. J. Controlled
Release, 110(1):67–78, 2005.
[136] F. Weinbreck, H. S. Rollema, R. H. Tromp, and C. G. De Kruif. Diffusivity of
whey protein and gum Arabic in their coacervates. Langmuir, 20(15):6389–6395,
2004.
[137] W. S. Price. Pulsed-field gradient nuclear magnetic resonance as a tool for
studying translational diffusion: Part I. Basic theory. Concepts Magn. Reson.,
9(5):299–336, 1997.
[138] W. S. Price. Pulsed-field gradient nuclear magnetic resonance as a tool for
studying translational diffusion: Part II. Experimental aspects. Concepts Magn.
Reson., 10(4):197–237, 1998.
[139] F. Brandl, M. Henke, S. Rothschenk, R. Gschwind, M. Breunig, T. Blunk, J. Teß-
mar, and A. Gopferich. Poly(ethylene glycol) based hydrogels for intraocular
applications. Adv. Eng. Mater., 9(12):1141–1149, 2007.
[140] P. J. Flory. Principles of polymer chemistry. Cornell University Press, Ithaca,
1953.
[141] J. C. Bray and E. W. Merrill. Poly(vinyl alcohol) hydrogels. Formation by
electron beam irradiation of aqueous solutions and subsequent crystallization.
J. Appl. Polym. Sci., 17(12):3779–3794, 1973.
144
Bibliography
[142] D. L. Elbert, A. B. Pratt, M. P. Lutolf, S. Halstenberg, and J. A. Hubbell.
Protein delivery from materials formed by self-selective conjugate addition
reactions. J. Controlled Release, 76(1-2):11–25, 2001.
[143] T. Canal and N. A. Peppas. Correlation between mesh size and equilibrium
degree of swelling of polymeric networks. J. Biomed. Mater. Res., 23(10):1183–
1193, 1989.
[144] G. P. Raeber, M. P. Lutolf, and J. A. Hubbell. Molecularly engineered PEG
hydrogels: a novel model system for proteolytically mediated cell migration.
Biophys. J., 89(2):1374–1388, 2005.
[145] L. G. Cima and S. T. Lopina. Network structures of radiation-crosslinked star
polymer gels. Macromolecules, 28(20):6787–6794, 1995.
[146] Y. Gnanou, G. Hild, and P. Rempp. Molecular structure and elastic behav-
ior of poly(ethylene oxide) networks swollen to equilibrium. Macromolecules,
20(7):1662–1671, 1987.
[147] D. M. Soumpasis. Theoretical analysis of fluorescence photobleaching recovery
experiments. Biophys. J., 41(1):95–97, 1983.
[148] C. Guse, S. Konnings, F. Kreye, F. Siepmann, A. Gopferich, and J. Siepmann.
Drug release from lipid-based implants: elucidation of the underlying mass
transport mechanisms. Int. J. Pharm., 314(2):137–144, 2006.
[149] G. M. Kavanagh and S. B. Ross-Murphy. Rheological characterisation of polymer
gels. Prog. Polym. Sci., 23(3):533–562, 1998.
[150] E. W. Merrill, K. A. Dennison, and C. Sung. Partitioning and diffusion of
solutes in hydrogels of poly(ethylene oxide). Biomaterials, 14(15):1117–1126,
1993.
[151] V. S. Sukumar and S. T. Lopina. Network model for the swelling properties
of end-linked linear and star poly(ethylene oxide) hydrogels. Macromolecules,
35(27):10189–10192, 2002.
145
Bibliography
[152] S. R. Lustig and N. A. Peppas. Solute diffusion in swollen membranes. IX.
Scaling laws for solute diffusion in gels. J. Appl. Polym. Sci., 36(4):735–747,
1988.
[153] M. Henke, F. Brandl, A. M. Gopferich, and J. K. Teßmar. Size dependent
release of fluorescent macromolecules and nanoparticles from radically cross-
linked hydrogels. Eur. J. Pharm. Biopharm., in press, 2009.
[154] N. A. Peppas, P. Bures, W. Leobandung, and H. Ichikawa. Hydrogels in
pharmaceutical formulations. Eur. J. Pharm. Biopharm., 50(1):27–46, 2000.
[155] J. W. DuBose, C. Cutshall, and A. T. Metters. Controlled release of teth-
ered molecules via engineered hydrogel degradation: Model development and
validation. J. Biomed. Mater. Res. A, 74A(1):104–116, 2005.
[156] H. Soyez, E. Schacht, M. Jelinkova, and B. Rihova. Biological evaluation of
mitomycin C bound to a biodegradable polymeric carrier. J. Controlled Release,
47(1):71–80, 1997.
[157] D. Seliktar, A. H. Zisch, M. P. Lutolf, J. L. Wrana, and J. A. Hubbell. MMP-2
sensitive, VEGF-bearing bioactive hydrogels for promotion of vascular healing.
J. Biomed. Mater. Res. A, 68A(4):704–716, 2004.
[158] R. G. Schoenmakers, P. van de Wetering, D. L. Elbert, and J. A. Hubbell.
The effect of the linker on the hydrolysis rate of drug-linked ester bonds. J.
Controlled Release, 95(2):291–300, 2004.
[159] S. Lee, R. B. Greenwald, J. McGuire, K. Yang, and C. Shi. Drug delivery
systems employing 1,6-elimination: Releasable poly(ethylene glycol) conjugates
of proteins. Bioconjug. Chem., 12(2):163–169, 2001.
[160] M. J. Roberts, M. D. Bentley, and J. M. Harris. Chemistry for peptide and
protein PEGylation. Adv. Drug Deliv. Rev., 54(4):459–476, 2002.
[161] R. B. Greenwald, K. Yang, H. Zhao, C. D. Conover, S. Lee, and D. Filpula.
Controlled release of proteins from their poly(ethylene glycol) conjugates: drug
146
Bibliography
delivery systems employing 1,6-elimination. Bioconjug. Chem., 14(2):395–403,
2003.
[162] D. Filpula and H. Zhao. Releasable PEGylation of proteins with customized
linkers. Adv. Drug Deliv. Rev., 60(1):29–49, 2008.
[163] M. M. Bradford. A rapid and sensitive method for the quantitation of micro-
gram quantities of protein utilizing the principle of protein-dye binding. Anal.
Biochem., 72(1-2):248–254, 1976.
[164] G. T. Hermanson. Bioconjugate Techniques. Academic Press, Amsterdam, The
Netherlands, 2nd edition, 2008.
[165] L. W. Dittert and T. Higuchi. Rates of hydrolysis of carbamate and carbonate
esters in alkaline solution. J. Pharm. Sci., 52:852–857, 1963.
[166] J. Hansen, N. Mrk, and H. Bundgaard. Phenyl carbamates of amino acids
as prodrug forms for protecting phenols against first-pass metabolism. Int. J.
Pharm., 81(2-3):253–261, 1992.
[167] E. Johnson, D. A. Berk, R. K. Jain, and W. M. Deen. Hindered diffusion in
agarose gels: test of effective medium model. Biophys. J., 70(2):1017–1023,
1996.
[168] S. C. De Smedt, A. Lauwers, J. Demeester, Y. Engelborghs, G. De Mey, and
M. Du. Structural information on hyaluronic acid solutions as studied by probe
diffusion experiments. Macromolecules, 27(1):141–146, 1994.
[169] A. Lucke, E. Fustella, J. Temar, A. Gazzaniga, and A. Gpferich. The effect
of poly(ethylene glycol)-poly(D,L-lactic acid) diblock copolymers on peptide
acylation. J. Controlled Release, 80(1-3):157–168, 2002.
[170] A. Lucke, J. Kiermaier, and A. Gpferich. Peptide acylation by poly(alpha-
hydroxy esters). Pharm. Res., 19(2):175–181, 2002.
[171] C. W. Patrick. Tissue engineering strategies for adipose tissue repair. Anat.
Rec., 263(4):361–366, 2001.
147
Bibliography
[172] B. Weiser, M. Neubauer, A. Gopferich, and T. Blunk. Encyclopedia of Biomate-
rials and Biomedical Engineering, volume 4, chapter Tissue Engineering, Fat,
pages 2725–2736. Marcel Dekker Inc., New York, 2 edition, 2005.
[173] B. Weiser, L. Prantl, T. E. O. Schubert, J. Zellner, C. Fischbach-Teschl, T. Spruß,
A. K. Seitz, J. Teßmar, A. Gopferich, and T. Blunk. In vivo development and
long-term survival of engineered adipose tissue depend on in vitro precultivation
strategy. Tissue Eng. Part A, 14(2):275–284, 2008.
[174] C. Fischbach, T. Spruß, B. Weiser, M. Neubauer, C. Becker, M. Hacker,
A. Gopferich, and T. Blunk. Generation of mature fat pads in vitro and
in vivo utilizing 3-D long-term culture of 3T3-L1 preadipocytes. Exp. Cell Res.,
300(1):54–64, 2004.
[175] A. Alhadlaq, M. Tang, and J. J. Mao. Engineered adipose tissue from human
mesenchymal stem cells maintains predefined shape and dimension: implications
in soft tissue augmentation and reconstruction. Tissue Eng., 11(3-4):556–566,
2005.
[176] P. N. Patel, C. K. Smith, and C. W. Patrick. Rheological and recovery properties
of poly(ethylene glycol) diacrylate hydrogels and human adipose tissue. J.
Biomed. Mater. Res. A, 73A(3):313–319, 2005.
[177] P. N. Patel, A. S. Gobin, J. L. West, and C. W. Patrick. Poly(ethylene glycol)
hydrogel system supports preadipocyte viability, adhesion, and proliferation.
Tissue Eng., 11(9-10):1498–1505, 2005.
[178] A. V. Vashi, E. Keramidaris, K. M. Abberton, W. A. Morrison, J. L. Wilson, A. J.
O’Connor, J. J. Cooper-White, and E. W. Thompson. Adipose differentiation
of bone marrow-derived mesenchymal stem cells using Pluronic F-127 hydrogel
in vitro. Biomaterials, 29(5):573–579, 2008.
[179] F. Brandl, F. Kastner, R. Gschwind, T. Blunk, J. Teßmar, and A. Gopferich.
Hydrogel-based drug delivery systems: Comparison of drug diffusivity and
release kinetics. J. Controlled Release, in press, 2009.
148
Bibliography
[180] M. P. Lutolf, G. P. Raeber, A. H. Zisch, N. Tirelli, and J. A. Hubbell. Cell-
Responsive Synthetic Hydrogels. Advanced Materials, 15(11):888–892, 2003.
[181] S. J. Bryant, C. R. Nuttelman, and K. S. Anseth. Cytocompatibility of UV and
visible light photoinitiating systems on cultured NIH/3T3 fibroblasts in vitro.
J. Biomater. Sci. Polym. Ed., 11(5):439–457, 2000.
[182] M. D. Timmer, H. Shin, R. A. Horch, C. G. Ambrose, and A. G. Mikos. In
vitro cytotoxicity of injectable and biodegradable poly(propylene fumarate)-
based networks: unreacted macromers, cross-linked networks, and degradation
products. Biomacromolecules, 4(4):1026–1033, 2003.
[183] C. Fischbach, J. Seufert, H. Staiger, M. Hacker, M. Neubauer, A. Gopferich,
and T. Blunk. Three-dimensional in vitro model of adipogenesis: comparison of
culture conditions. Tissue Eng., 10(1-2):215–229, 2004.
[184] M. Mutter, R. Uhmann, and E. Bayer. Die Liquid-Phase-Methode zur Pep-
tidsynthese; Strategie und experimentelle Grundlagen. Justus Liebigs Ann.
Chem., 1975(5):901–915, 1975.
[185] S. Udenfriend, S. Stein, P. Bohlen, W. Dairman, W. Leimgruber, and M. Weigele.
Fluorescamine: A reagent for assay of amino acids, peptides, proteins, and
primary amines in the picomole range. Science, 178(4063):871–872, 1972.
[186] Y.-J. Kim, R. L. Y. Sah, J.-Y. H. Doong, and A. J. Grodzinsky. Fluorometric
assay of DNA in cartilage explants using Hoechst 33258. Anal. Biochem.,
174(1):168–176, 1988.
[187] J. L. Ramırez-Zacarıas, F. Castro-Munozledo, and W. Kuri-Harcuch. Quantita-
tion of adipose conversion and triglycerides by staining intracytoplasmic lipids
with Oil red O. Histochem. Cell Biol., 97(6):493–497, 1992.
[188] Y. Teramura, Y. Kaneda, T. Totani, and H. Iwata. Behavior of synthetic
polymers immobilized on a cell membrane. Biomaterials, 29(10):1345–1355,
2008.
149
Bibliography
[189] A. Nordwig and L. Strauch. Stabilitatsverhalten der Kollagenase aus Clostridium
histolyticum. Hoppe-Seyler’s Z. Physiol. Chem., 330:153–160, 1963.
[190] J. L. West and J. A. Hubbell. Polymeric biomaterials with degradation sites for
proteases involved in cell migration. Macromolecules, 32(1):241–244, 1999.
[191] B. K. Mann, A. S. Gobin, A. T. Tsai, R. H. Schmedlen, and J. L. West.
Smooth muscle cell growth in photopolymerized hydrogels with cell adhesive
and proteolytically degradable domains: synthetic ECM analogs for tissue
engineering. Biomaterials, 22(22):3045–3051, 2001.
[192] M. D. Sternlicht and Z. Werb. How matrix metalloproteinases regulate cell
behavior. Annu. Rev. Cell Dev. Biol., 17:463–516, 2001.
[193] B. M. Spiegelman and C. A. Ginty. Fibronectin modulation of cell shape and
lipogenic gene expression in 3T3-adipocytes. Cell, 35(3, Part 2):657–666, 1983.
[194] K. C. O’Connor, H. Song, N. Rosenzweig, and D. A. Jansen. Extracellular
matrix substrata alter adipocyte yield and lipogenesis in primary cultures of
stromal-vascular cells from human adipose. Biotechnol. Lett., 25(23):1967–1972,
2003.
[195] C. M. Smas and H. S. Sul. Control of adipocyte differentiation. Biochem. J.,
309(3):697–710, 1995.
[196] G. J. Hausman, J. T. Wright, and R. L. Richardson. The influence of extracellular
matrix substrata on preadipocyte development in serum-free cultures of stromal-
vascular cells. J. Anim. Sci., 74(9):2117–2128, 1996.
[197] C. W. Patrick and X. Wu. Integrin-mediated preadipocyte adhesion and
migration on laminin-1. Ann. Biomed. Eng., 31(5):505–514, 2003.
[198] B. Jeong, S. W. Kim, and Y. H. Bae. Thermosensitive sol-gel reversible hydrogels.
Adv. Drug Deliv. Rev., 54(1):37–51, 2002.
[199] C. Framme, F. Brandl, S. Rothschenk, A. Gopferich, H. Helbig, and K. Kobuch.
Development and evaluation of new hydrogels for vitreous substitution and
intravitreal drug application. 106th DOG Congress, Berlin, Germany, 2008.
150
Bibliography
[200] F. Brandl, K. Kobuch, S. Rothschenk, T. Blunk, J. Teßmar, and A. Gopferich.
In situ-forming hydrogels for intraocular drug delivery. 8th World Biomaterials
Congress, Amsterdam, The Netherlands, 2008.
[201] K. Kobuch, W. A. Herrmann, C. Framme, H. G. Sachs, V.-P. Gabel, and J. Hil-
lenkamp. Maintenance of adult porcine retina and retinal pigment epithelium
in perfusion culture: Characterisation of an organotypic in vitro model. Exp.
Eye Res., 86(4):661–668, 2008.
151
List of Figures
1.1 Principle of in situ forming hydrogels . . . . . . . . . . . . . . . . . . 7
2.1 Repositioning of cells by mechanotaxis on surfaces with micropatterned
Young’s modulus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16
2.2 Schematic diagram of atomic force microscopy (AFM) . . . . . . . . . 21
2.3 Schematic diagram of magnetic resonance elastography (MRE) . . . . 22
2.4 Principle of fluorescence resonance energy transfer (FRET) . . . . . . 24
2.5 Michael-type addition reaction between vinylsulfone-functionalized
PEG macromers and cysteine containing peptides . . . . . . . . . . . 26
2.6 Schematic illustration of cell-responsive hydrogels . . . . . . . . . . . 33
3.1 Synthesis of PEG2k-NH2 and 4armPEG10k-SPA . . . . . . . . . . . . 43
3.2 Typical rheogram of a gel formed by polymerization of PEG2k-NH2
with 4armPEG10k-SPA . . . . . . . . . . . . . . . . . . . . . . . . . . 48
3.3 Dependence of gelation time and gel strength on the stoichiometric
ratio, polymer concentration, pH, and temperature . . . . . . . . . . 49
3.4 1H-NMR spectra of a hydrogel formed by reaction of 4armPEG10k-SPA
and PEG2k-NH2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53
3.5 Cell viability after exposure to hydrogel extracts . . . . . . . . . . . . 54
3.6 Release of FITC-dextrans from non-degradable hydrogels . . . . . . . 55
4.1 Image stack of a typical FRAP experiment . . . . . . . . . . . . . . . 71
4.2 Diffusion coefficients of FITC-dextrans determined by FRAP . . . . . 73
153
List of Figures
4.3 Release of FITC-dextrans from non-degradable hydrogels . . . . . . . 75
5.1 Schematic illustration of hydrolytically degradable hydrogels . . . . . 80
5.2 Polymers derived from branched poly(ethylene glycol) . . . . . . . . . 83
5.3 Typical rheograms of degradable and non-degradable hydrogels . . . . 92
5.4 Degradation of hydrogels . . . . . . . . . . . . . . . . . . . . . . . . . 94
5.5 Mobile fractions of FITC-dextran and FITC-BSA . . . . . . . . . . . 96
5.6 Release of FITC-BSA and lysozyme from biodegradable hydrogels . . 98
6.1 Schematic illustration of injectable, biointeractive hydrogels for soft
tissue engineering applications . . . . . . . . . . . . . . . . . . . . . . 105
6.2 Degradation of hydrogels . . . . . . . . . . . . . . . . . . . . . . . . . 117
6.3 Comparison of 2-D and 3-D cell culture . . . . . . . . . . . . . . . . . 120
6.4 3T3-L1 adipocytes cultured in hydrogels without and with 1 µmol/mL
YIGSR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122
6.5 Long-term culture of 3T3-L1 adipocytes . . . . . . . . . . . . . . . . 123
7.1 Polymers derived from four-armed poly(ethylene glycol) . . . . . . . . 127
7.2 PEG hydrogel formed by reaction between branched PEG-succinimidyl
propionates and branched PEG-amines. . . . . . . . . . . . . . . . . . 128
7.3 PEG hydrogels in perfusion organ culture . . . . . . . . . . . . . . . . 130
154
List of Tables
2.1 Mechanical properties of different biological tissues . . . . . . . . . . 19
3.1 Composition of hydrogels and conditions of gel preparation . . . . . . 45
4.1 Physicochemical characteristics of the prepared hydrogels . . . . . . . 68
4.2 Hydrodynamic radii and diffusion coefficients of FITC-dextrans . . . 70
4.3 Mobile fractions and diffusion coefficients of FITC-dextrans . . . . . . 72
5.1 Composition of the prepared hydrogels . . . . . . . . . . . . . . . . . 87
5.2 Physicochemical characteritsics of the prepared hydrogels . . . . . . . 93
6.1 Physicochemical characteristics of the prepared hydrogels . . . . . . . 115
155
Appendix
157
Acronyms
2-D . . . . . . . . . . . . two-dimensional
3-D . . . . . . . . . . . . three-dimensional
AFM . . . . . . . . . . . atomic force microscopy
Ala . . . . . . . . . . . . alanine
AMD . . . . . . . . . . . age-related macular degeneration
ANOVA . . . . . . . . . analysis of variance
Arg . . . . . . . . . . . . arginine
Asp . . . . . . . . . . . . aspartic acid
BMP-2 . . . . . . . . . . bone morphogenic protein-2
BMSC . . . . . . . . . . bone marrow stromal cell
BSA . . . . . . . . . . . . bovine serum albumin13C-NMR . . . . . . . . carbon-13 nuclear magnetic resonance
Cys . . . . . . . . . . . . cysteine
DCC . . . . . . . . . . . N,N’ -dicyclohexylcarbodiimide
DCM . . . . . . . . . . . methylene chloride
DCU . . . . . . . . . . . N,N’ -dicyclohexylurea
DIAD . . . . . . . . . . . diisopropyl azodicarboxylate
DIPEA . . . . . . . . . . N,N’ -diisopropylethylamine
DMA . . . . . . . . . . . dynamic mechanical analysis
DMEM . . . . . . . . . . Dulbecco’s modified Eagle’s medium
DNA . . . . . . . . . . . desoxyribonucleic acid
DOSY . . . . . . . . . . diffusion ordered NMR spectroscopy
DRG . . . . . . . . . . . dorsal root ganglion
159
Acronyms
DSC . . . . . . . . . . . . N,N’ -discuccinimidyl carbonate
ECM . . . . . . . . . . . extracellular matrix
EDTA . . . . . . . . . . ethylenediaminetetraacetic acid
FBS . . . . . . . . . . . . fetal bovine serum
FITC . . . . . . . . . . . fluoresceine isothiocyanate
FRAP . . . . . . . . . . fluorescence recovery after photobleaching
FRET . . . . . . . . . . . fluorescence resonance energy transfer
G . . . . . . . . . . . . . guluronic acid
GAG . . . . . . . . . . . glycosaminoglycan
Gly . . . . . . . . . . . . glycine1H-NMR . . . . . . . . . proton nuclear magnetic resonance
HBOEC . . . . . . . . . human blood outgrowth endothelial cell
HEPES . . . . . . . . . . 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
HMBC . . . . . . . . . . heteronuclear multiple bond coherence
HOSu . . . . . . . . . . . N-hydroxysuccinimide
HUVEC . . . . . . . . . human umbilical vein endothelial cell
IBMX . . . . . . . . . . . 3-isobutyl-methylxanthine
Ile . . . . . . . . . . . . . isoleucine
Leu . . . . . . . . . . . . leucine
Lys . . . . . . . . . . . . lysine
M . . . . . . . . . . . . . mannuronic acid
α-MEM . . . . . . . . . . Minimum Essential Medium alpha modification
µCT . . . . . . . . . . . . microcomputed tomography
µMRE . . . . . . . . . . microscopic magnetic resonance elastography
MMP . . . . . . . . . . . matrix metalloproteinase
MR . . . . . . . . . . . . magnetic resonance
MRE . . . . . . . . . . . magnetic resonance elastography
MRI . . . . . . . . . . . . magnetic resonance imaging
MW . . . . . . . . . . . . molecular weight
NHS . . . . . . . . . . . . N-hydroxysuccinimide
NMR . . . . . . . . . . . nuclear magnetic resonance
PAA . . . . . . . . . . . . poly(acrylic acid)
160
Acronyms
PAAM . . . . . . . . . . polyacrylamide
PBS . . . . . . . . . . . . phosphate buffered saline
PCR . . . . . . . . . . . polymerase chain reaction
PDMS . . . . . . . . . . poly(dimethylsiloxane)
PDR . . . . . . . . . . . proliferative diabetic retinopathy
PEG . . . . . . . . . . . poly(ethylene glycol)
PEG2k-OH . . . . . . . poly(ethylene glycol), molecular weight 2 kDa
4armPEG10k-OH . . . four-armed poly(ethylene glycol), molecular weight 10 kDa
PGA . . . . . . . . . . . poly(glycolic acid)
PLA . . . . . . . . . . . . poly(lactic acid)
PLGA . . . . . . . . . . poly(lactide-co-glycolide)
PPh3 . . . . . . . . . . . triphenylphosphine
Pro . . . . . . . . . . . . proline
PSPD . . . . . . . . . . . position sensitive photodiode
PVA . . . . . . . . . . . . poly(vinyl alcohol)
RF . . . . . . . . . . . . . radiofrequency
RGD . . . . . . . . . . . arginine–glycine–aspartic acid
RGDSP . . . . . . . . . . arginine–glycine–aspartic acid–serine–proline
SDS . . . . . . . . . . . . sodium dodecyl sulfate
Ser . . . . . . . . . . . . . serine
SMC . . . . . . . . . . . smooth muscle cell
TGF-β3 . . . . . . . . . transforming growth factor-β3
THF . . . . . . . . . . . tetrahydrofuran
TLC . . . . . . . . . . . . thin layer chromatography
Tris . . . . . . . . . . . . tris(hydroxymethyl)aminomethane
Tyr . . . . . . . . . . . . tyrosine
UV . . . . . . . . . . . . ultraviolet
VEGF . . . . . . . . . . vascular endothelial growth factor
YIGSR . . . . . . . . . . tyrosine–isoleucine–glycine–serine–arginine
161
Symbols
c . . . . . . . . . . . . . . concentration
χ1 . . . . . . . . . . . . . Flory-Huggins interaction parameter
Cn . . . . . . . . . . . . . Flory characteristic ratio
D . . . . . . . . . . . . . diffusion coefficient
D0 . . . . . . . . . . . . . theoretical diffusion coefficient in water
∆ . . . . . . . . . . . . . diffusion time
δ . . . . . . . . . . . . . . chemical shift
δ . . . . . . . . . . . . . . gradient pulse length
Dg . . . . . . . . . . . . . estimated diffusion coefficient in hydrogels
η . . . . . . . . . . . . . . dynamic viscosity
f . . . . . . . . . . . . . . functionality of the cross-link
f(t) . . . . . . . . . . . . normalized fluorescence intensity
G∗ . . . . . . . . . . . . . complex shear modulus
G′ . . . . . . . . . . . . . storage modulus
G′′ . . . . . . . . . . . . . loss modulus
γ∗ . . . . . . . . . . . . . oscillatory shear strain
Ge . . . . . . . . . . . . . equilibrium modulus
h . . . . . . . . . . . . . . height of the gel cylinder
I0 . . . . . . . . . . . . . modified Bessel function of the first kind of zero order
I1 . . . . . . . . . . . . . modified Bessel function of the first kind of first order
k . . . . . . . . . . . . . . mobile fraction
k . . . . . . . . . . . . . . Boltzmann constant
l . . . . . . . . . . . . . . bond length along the polymer backbone
163
Symbols
λem . . . . . . . . . . . . emission wavelength
λex . . . . . . . . . . . . . excitation wavelength
Mc . . . . . . . . . . . . . average molecular weight between cross-links
mp . . . . . . . . . . . . . total mass of polymer in the hydrogel
Mr . . . . . . . . . . . . . molecular mass of the polymer repeating unit
Mt . . . . . . . . . . . . . absolute cumulative amount released at time point t
µe . . . . . . . . . . . . . number of moles of cross-links
M∞ . . . . . . . . . . . . absolute cumulative amount released at infinity
νe . . . . . . . . . . . . . number of moles of elastically active chains
Q . . . . . . . . . . . . . volumetric swelling ratio
qn . . . . . . . . . . . . . roots of the Bessel function of the first kind of zero order
R . . . . . . . . . . . . . molar gas constant
r . . . . . . . . . . . . . . stoichiometric ratio
r . . . . . . . . . . . . . . radius of the gel cylinder
rH . . . . . . . . . . . . . hydrodynamic radius
σ∗ . . . . . . . . . . . . . oscillatory shear stress
T . . . . . . . . . . . . . absolute temperature
t . . . . . . . . . . . . . . time
τD . . . . . . . . . . . . . characteristic diffusion time
V1 . . . . . . . . . . . . . molar volume of the solvent
v2c . . . . . . . . . . . . . polymer fraction of the gel after cross-linking
v2s . . . . . . . . . . . . . polymer fraction of the gel in the swollen state
Vgc . . . . . . . . . . . . . gel volume after cross-linking
Vgs . . . . . . . . . . . . . gel volume after swelling
Vp . . . . . . . . . . . . . volume of the dry polymer
w . . . . . . . . . . . . . radius of the bleached spot
ξ . . . . . . . . . . . . . . average network mesh size
Y . . . . . . . . . . . . . ratio of the critical volume
164
Curriculum vitae
Personal information
Name: Ferdinand Paul Brandl
Date of birth: 02/17/1979
Place of birth: Hirschau, Germany
Nationality: German
Marital Status: Single
Professional training
Since 12/2004 University of Regensburg
PhD program at the Department of Pharmaceutical
Technology (Prof. Dr. Achim Gopferich)
12/17/2004 Approbation zum Apotheker
(Acquisition of the license to practice as pharmacist)
05/2004 – 10/2004 Dispensary of the University Hospital, Regensburg
Practical training
165
Curriculum vitae
11/2003 – 04/2004 Rosen Apotheke, Amberg
Practical training
11/1999 – 09/2003 University of Regensburg
Pharmaceutics course
Civilian service
07/1998 - 07/1999 Bund Naturschutz in Bayern e.V., Wiesenfelden
(Environmental association of Bavaria)
Education
06/26/1998 Allgemeine Hochschulreife
(General qualification for university entrance)
09/1989 - 06/1998 Erasmus-Gymnasium, Amberg
(Grammar School)
09/1985 - 07/1989 Albert-Schweitzer-Schule, Amberg
(Primary School)
166
List of publications
Publications in scientific journals
1. Ferdinand Brandl, A. Seitz, J. Teßmar, T. Blunk, and A. Gopferich. Biointeractive hydrogelsfor adipose tissue engineering. Submitted to Biomaterials. (Chapter 6)
2. Ferdinand Brandl, J. Teßmar, T. Blunk, and A. Gopferich. Biodegradable hydrogels fortime-controlled release of tethered peptides or proteins. Submitted to Biomacromolecules.(Chapter 5)
3. Ferdinand Brandl, F. Kastner, R. Gschwind, T. Blunk, J. Teßmar, and A. Gopferich.Hydrogel-based drug delivery systems: Comparison of drug diffusivity and release kinetics. J.Controlled Release, in press. (Chapter 4)
4. M. Henke, Ferdinand Brandl, A. Gopferich, and J. Teßmar. Size dependent release offluorescent macromolecules and nanoparticles from radically cross-linked hydrogels. Eur. J.Pharm. Biopharm., in press.
5. F. Sommer, Ferdinand Brandl, B. Weiser, J. Teßmar, T. Blunk, and A. Gopferich. FACSas useful tool to study distinct hyalocyte populations. Exp. Eye Res. 88(5):995–999, 2009.
6. F. Sommer, K. Pollinger, Ferdinand Brandl, B. Weiser, J. Teßmar, T. Blunk, and A.Gopferich. Hyalocyte proliferation and ECM accumulation modulated by bFGF and TGF-β1.Graefes Arch. Clin. Exp. Opthalmol. 246(9):1275–1284, 2008.
7. Ferdinand Brandl, M. Henke, S. Rothschenk, R. Gschwind, M. Breunig, T. Blunk, J. Teßmar,and A. Gopferich. Poly(ethylene glycol) based hydrogels for intraocular applications. Adv.Eng. Mater. 9(12):1141–1149, 2007. (Chapter 3)
167
List of publications
8. F. Sommer, K. Kobuch, Ferdinand Brandl, B. Wild, C. Framme, B. Weiser, J. Teßmar,V.-P. Gabel, T. Blunk, and A. Gopferich. Ascorbic acid modulates proliferation and extracel-lular matrix accumulation of hyalocytes. Tissue Eng. 13(6):1281–1289, 2007.
9. Ferdinand Brandl, F. Sommer, and A. Gopferich. Rational design of hydrogels for tissueengineering: Impact of physical factors on cell behaviour. Biomaterials 28(2):134-146, 2007.(Chapter 2)
10. D. Eyrich, Ferdinand Brandl, B. Appel, H. Wiese, G. Maier, M. Wenzel, R. Staudenmaier,A. Gopferich, and T. Blunk. Long-term stable fibrin gels for cartilage engineering. Biomaterials28(1):55–65, 2007.
Book chapters
1. J. K. Teßmar, Ferdinand Brandl, and A. M. Gopferich. Fundamentals of Tissue Engineeringand Regenerative Medicine, chapter Hydrogels for Tissue Engineering, pages 495–517. Springer,Berlin, 2009.
2. F. Sommer, Ferdinand Brandl, and A. Gopferich. Advances in Experimental Medicine andBiology Vol. 585, chapter Ocular Tissue Engineering, pages 413–429. Springer, Berlin, 2007.
Conference abstracts
1. Ferdinand Brandl, T. Blunk, J. Teßmar, and A. Gopferich. Novel biodegradable hydrogelsfor peptide and protein delivery. 2nd PharmSciFair, Nice, France (2009).
2. Ferdinand Brandl, K. Kobuch, S. Rothschenk, T. Blunk, J. Teßmar, and A. Gopferich.In situ-gelling hydrogels for intraocular drug delivery. 8th World Biomaterials Congress,Amsterdam, The Netherlands (2008).
3. A. Gopferich, Ferdinand Brandl, and R. Knerr. Tools for the characterization of biomimeticand interactive polymers. 8th World Biomaterials Congress, Amsterdam, Netherlands (2008).
4. Ferdinand Brandl, S. Rothschenk, A. Blaimer, T. Blunk, J. Teßmar, and A. Gopferich. Insitu-forming hydrogels for intraocular drug delivery. 6th World Meeting on Pharmaceutics,Biopharmaceutics and Pharmaceutical Technology, Barcelona, Spain (2008).
168
5. S. Rothschenk, Ferdinand Brandl, J. Teßmar, T. Blunk, and A. Gopferich. Release ofimmunoglobulin G from in situ-forming hydrogels for intraocular drug delivery. 6th WorldMeeting on Pharmaceutics, Biopharmaceutics and Pharmaceutical Technology, Barcelona,Spain (2008).
6. C. Framme, Ferdinand Brandl, S. Rothschenk, A. Gopferich, H. Helbig, and K. Kobuch.Development and evaluation of new hydrogels for vitreous substitution and intravitreal drugapplication. 106th DOG Congress, Berlin, Germany (2008).
7. A. Gopferich, Ferdinand Brandl, J. Teßmar, and M. Breunig. Strategies for local pro-tein and nucleic acid delivery. 3rd International Conference on Tissue Engineering, Rhode,Greece (2008).
8. Ferdinand Brandl, S. Rothschenk, M. Breunig, J. Teßmar, T. Blunk, and A. Gopferich.Polyethylene glycol macromers as building blocks for versatile hydrogels. 1st InternationalCongress on Biohydrogels, Viareggio, Italy (2007).
9. Ferdinand Brandl, F. Launay, F. Sommer, T. Blunk, J. Teßmar, and A. Gopferich. Insitu-gelling poly(ethylene glycol) based hydrogels for biomedical applications. PolyPharma,Halle (Saale), Germany (2006).
10. Ferdinand Brandl, F. Sommer, U. Lungwitz, T. Blunk, J. Teßmar, and A. Gopferich. Insitu-gelling hydrogels based on poly(ethylene glycol). 2nd International Symposium InterfaceBiology of Implants, Rostock, Germany (2006).
11. F. Sommer, Ferdinand Brandl, B. Weiser, J. Teßmar, T. Blunk, and A. Gopferich. Hyalo-cytes within the vitreous body – a homogenous population? First evidence for two distinctpopulations. 2nd International Conference on Strategies in Tissue Engineering, Wurzburg,Germany (2006).
12. F. Sommer, K. Pollinger, Ferdinand Brandl, B. Weiser, J. Teßmar, T. Blunk, and A.Gopferich. Towards a cell-based vitreous substitute – The effect of bFGF and TGF-β1 onhyalocytes. 33rd Annual Meeting & Exposition of the Controlled Release Society, Vienna,Austria (2006).
13. F. Sommer, K. Kobuch, Ferdinand Brandl, B. Wild, B. Weiser, V.-P. Gabel, T. Blunk,and A. Gopferich. Ascorbic acid for in vitro hyalocyte culture – an important factor towardsa cellular vitreous substitute. 2nd International Conference on Tissue Engineering, Crete,Greece (2005).
14. F. Sommer, K. Kobuch, Ferdinand Brandl, B. Wild, B. Weiser, V.-P. Gabel, T. Blunk,and A. Gopferich. Ascorbic acid influences hyalocytes on the molecular level – increased
169
List of publications
expression of collagen type V/XI. 4th Annual Meeting of the European Tissue EngineeringSociety, Munich, Germany (2005).
170
Acknowledgments
An dieser Stelle mochte ich all jenen meinen herzlichen Dank aussprechen, die zum
Gelingen dieser Arbeit und der unvergesslichen Zeit am Lehrstuhl beigetragen haben.
Mein ganz besonderer Dank gilt Herrn Prof. Dr. Achim Gopferich fur die Uberlassung
des Themas, seine kontinuierliche Unterstutzung bei dessen Bearbeitung, sein stetes
Interesse am Fortgang der Experimente und fur die Freiheit, eigene Ideen und Ansatze
verwirklichen zu konnen. Die zahlreichen wissenschaftlichen Diskussion und Anre-
gungen waren immer sehr hilfreich fur mich. Ganz besonders mochte ich mich auch
fur die Moglichkeit bedanken, meine Arbeiten auf nationalen und internationalen
Kongressen zu prasentieren.
Herrn Dr. Torsten Blunk und Herrn Dr. Jorg Teßmar danke ich sehr herzlich fur
unzahlige konstruktive Ratschlage, zahlreiche wissenschaftliche Diskussionen und die
freundschaftliche Zusammenarbeit. Insbesondere mochte ich ihnen fur die Unter-
stutzung bei der Niederschrift dieser Arbeit danken.
Mein Dank gilt auch Frau Dr. Karin Kobuch, Herrn PD Dr. Carsten Framme, Brigitte
Wild und Petra Eberl fur die gute Zusammenarbeit im Glaskorper-Projekt.
Fur die finanzielle Unterstutzung des Projektes mochte ich mich bei der Bayerischen
Forschungsstiftung bedanken (AZ 616/04).
171
Acknowledgments
Weiterhin danke ich allen derzeitigen und ehemaligen Kolleginnen und Kollegen
am Lehrstuhl fur das gute, oft freundschaftliche Arbeitsklima, die konstruktive
Zusammenarbeit und die vielen außerfachlichen Gesprache und Aktivitaten.
Mein herzlicher Dank gilt insbesondere:
• Dr. Florian Sommer und Dr. Stefan Rothschenk fur die gute und motivierte
Zusammenarbeit im Glaskorper-Projekt – in guten wie in schlechten Zeiten
• Annina Seitz, Renate Liebl und Julia Baumer fur die Durchfuhrung von zahl-
reichen Zellkulturexperimenten
• Dr. Thomas Burgemeister, Fritz Kastner, Annette Schramm und Georgine
Stuhler fur die Durchfuhrung zahlloser NMR-Messungen
• Nadine Hammer und Alexandra Frimberger fur die Unterstutzung im Labor
• Andrea Blaimer fur die Unterstutzung am Rheometer
• Angelika Berie, Lydia Frommer, Stefan Kolb, Liane Ottl und Edith Schindler
fur vielerlei technische und organisatorische Hilfe
• Leon Bellan, Allison Dennis, Leda Klouda und Peter Yang fur die schnelle und
gewissenhafte Durchsicht zahlreicher Manuskripte einschließlich dieser Arbeit
• Dr. Bernhard Appel, Axel Ehmer, Matthias Henke, Constantin Hozsa und Dr.
Florian Sommer fur die engagierte Zusammenarbeit im bitsandbytes-Team
• Constantin Hozsa fur die heitere Laborgemeinschaft und seine Freundschaft
• Dr. Bernhard Appel, Christian Becker, Dr. Uta Lungwitz, Dr. Angelika Maschke,
Dr. Barbara Weiser und allen anderen Doktoranden der alten Riege fur die
freundliche Aufnahme und die vielen gemeinsamen Unternehmungen
Besonderer Dank gilt meinen Eltern, die mir diesen Weg ermoglicht haben und mich –
wo immer sie nur konnten – bestarkt und unterstutzt haben.
172
Dieses Dokument wurde mit LATEX 2ε erstellt.
KOMA-Script (2009/04/03 v3.03a KOMA-Script)
173